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Full text of "Buzzards Bay caged mussel pilot biomonitoring study, 1987-1988"

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PILOT BIOMONITORING STUDY 

1987-1988 




I 



XL? 11 *** ^^. 



Massachusetts Executive Office of Environmental Affairs 
John P. DeVillars, Secretary 

Department of Environmental Protection 
Daniel S. Greenbaum, Commissioner 




Division of Water Pollution Control 
Brian M. Donahoe, Director 



315DbbDltit l DS4fi i: ] 



BUZZARDS BAY 

CAGED MUSSEL PILOT BIOMONITORING STUDY 

1987 - 1988 



Prepared by 

Christine L. Duerring 
Environmental Analyst 



MASSACHUSETTS DEPARTMENT OF ENVIRONMENTAL PROTECTION 

DIVISION OF WATER POLLUTION CONTROL 

TECHNICAL SERVICES BRANCH 

WESTBOROUGH, MASSACHUSETTS 

FOR 

U.S. ENVIRONMENTAL PROTECTION AGENCY 

REGION I 

WATER MANAGEMENT DIVISION 

BOSTON, MASSACHUSETTS 



Massachusetts Executive Office of Environmental Affairs 
John P. DeVillars, Secretary 

Department of Environmental Protection 
Daniel S. Greenbaum, Commissioner 

Division of Water Pollution Control 
Brian M. Donahoe, Director 



OCTOBER 1990 



Publication No. 1 6 , 500- 1 07-37- 1 1-90-CR 

Approved by: Ric Murphy, State Purchasing Agent 



TITLE: 



Buzzards Bay Caged Mussel Pilot Biomonitoring Study 



DATE: 



October, 1990 



AUTHOR : 



Christine L. Duerring, Environmental Analyst III 



REVIEWED BY: 




Steven G. Halterman 
Environmental Engineer 



APPROVED BY: 




,/ 



u ? y (^Yj^ br^ 4 * 



Alan N. Coopermapr, P.E. 
Supervisor, Technical Services Branch 



ii 



FOREWORD 



The Division of Water Pollution Control in 1987 proposed and received funding 
from the U.S. Environmental Protection Agency (EPA) to conduct a pilot 
biomonitoring study in Buzzards Bay using caged mussels ( Mytilus edulis ) . The 
study is part of a national estuarine management program developed by the U.S. 
EPA Office of Marine and Estuarine Protection and Region I of the EPA for 
Buzzards Bay. The program was initiated to promote and develop coordinated 
efforts between federal, state, local authorities, research institutions and the 
public to identify, correct, and monitor environmental problems affecting this 
nation's estuaries. 



111 



ACKNOWLEDGMENTS 



The following people and groups are gratefully acknowledged for their assistance 
in conducting various aspects of this study: 

Patricia Austin, Lawrence Gil, Steve Halterman, Jay Loubris, Steve McGee, Cathy 
O'Riordan, crew and captain of the fishing boat Phalarope, and Gerry Szal for 
assisting in the collection and/or deployment of the mussels while maintaining 
a sense of humor in the face of (usually) adverse weather conditions; 

Brad Burke, New Bedford Shellfish Constable and his assistant David Goulart, who 
provided boat transportation and muscle in deploying and collecting the cages 
throughout the entire study. Their reliable assistance and knowledge of the area 
eased the field work effort considerably; 

Staff of the Lawrence Experiment Station, notably Ken Hulme, Rosario Grasso and 
Alba Flaherty for performing the special water quality and tissue analyses; 

Ken Hulme together with Nina Dustin and Jack Schwartz of the Division of Marine 
Fisheries, Cat Cove Laboratory also offered helpful comments concerning inter- 
laboratory calibration exercises; 

Bruce Tripp who offered encouragement and technical advice during the study; 

Judy Capuzzo and Donald Phelps for helpful review of the data; 

And lastly, but most important, this study would not have been possible without 
the use of the laboratory facilities at the EPA Environmental Research Laboratory 
in Narragansett, RI and the generous assistance and advice of Skip Nelson in 
designing and implementing the study. 



TABLE OF CONTENTS 

TITLE PAGE 

Acknowledgments v 

Abstract ix 

List of Tables xi 

List of Figures xiii 

Introduction 1 

Methods and Description of Study Site 5 

Results 14 

Discussion 37 

Summary 45 

Bibliography 46 

Appendix A: Field and LES Laboratory Methodology 49 

Appendix B: Field and Laboratory Data 73 

Appendix C: Sample Statistical Calculations 85 

Appendix D: Division of Marine Fisheries Project 89 
Plan and Laboratory Methodology 



VII 



Digitized by the Internet Archive 

in 2013 with funding from 

Boston Library Consortium Member Libraries 



http://archive.org/details/buzzardsbaycageOOduer 



ABSTRACT 



Buzzards Bay Caged Mussel Pilot Biomonitoring Study 1987 - 1988 

A caged mussel pilot biomonitoring study was conducted in Clarks Cove, New 
Bedford/Dartmouth, Massachusetts from October 1987 to September 1988. Mussels 
were deployed at three stations for five consecutive, 60-day exposure periods. 
Mussel tissue was analyzed for the trace elements: As, Cd, Cr, Cu, Hg, Ni, Pb, 
Zn, as well as total and fecal coliform bacteria and polychlorinated biphenyls 
(PCBs), and percent lipid content before and after the exposure periods. 

Trace element tissue concentration was extremely variable at all of the stations. 
Within station (replicate) variability was usually high and masked between 
station differences in trace element concentration for many of the deployments. 
However, significant differences were detected between baseline and one or more 
of the Clarks Cove Stations for tissue concentrations of arsenic, zinc, and lead 
for several of the exposure periods. None of the Clarks Cove Stations (A, B, 
or C) exhibited significant differences in trace element tissue concentration 
from each other, indicating bio-available trace element concentration was not 
spatially different in Clarks Cove. 

Bacteria concentration in the mussel tissue was variable and showed no consistent 
pattern throughout the study. Based on these results this technique is not 
recommended for long-term monitoring of coliform densities in coastal areas. 

PCB tissue concentration between baseline and Clarks Cove Stations showed a 
consistent pattern of low baseline values, highest concentration at Station A, 
next highest at Station B, and low at Station C, indicating that this method may 
be effective for monitoring PCB concentration in coastal areas. 

Inter-laboratory calibration exercises performed between the Lawrence Experiment 
Station and the Division of Marine Fisheries, Cat Cove Laboratory showed large 
inter-laboratory differences in results from mussel tissue analyzed for trace 
element concentration from the Clarks Cove study sites. However, results from 
similar analyses of EPA prepared standard "mega mussel" samples showed good 
inter-laboratory agreement. 



ix 



LIST OF TABLES 

TABLE TITLE PAGE 

1 Clarks Cove Combined Sewer Overflows 12 

2 Clarks Cove Sediment Data: Trace Elements, PCBs, PAHs, 16 
Percent Total Volatile Solids 

3 Percent Mortality of Mussels and Cage Loss 18 

4 Mussel Tissue Total and Fecal Coliform Densities 20 

5 Summary of Kruskal-Wallis Nonparametric Analysis of 30 
Variance 

6 Summary of Tukey-type Nonparametric Multiple Comparison 31 
Tests 

7 Inter-laboratory Calibration Results - Trace Element 34 
Concentrations 

8 Inter-laboratory Calibration Results - Standard 36 
"Mega Mussel" Tissue 

9 Tissue Trace Element Concentration - Comparison to Other 41 
Studies 

A-l Common Sample Treatment Methods 50 

A-2 Parameter and Collection Methods Employed at Sediment 51 
Stations 

A-3 Summary of Rated Accuracy of Field Meters and Unit of 52 
Measure 

A-4 Parameters and Analytical Methods for Water Samples 53 

A-5 Parameters and Analytical Methods for Sediment Samples 55 

A-6 Parameters and Analytical Methods for Tissue Samples 57 

A-7 Method for Chlorophyll a Analysis 59 

B-l Clarks Cove Water Quality Data - Field Measurements 74 

B-2 Clarks Cove Water Quality Data - Chemical Parameters 77 

B-3 Results of Mussel Tissue Trace Element Analysis 80 

B-4 Mussel Tissue PCB Concentrations 83 

B-5 Percent Lipid Concentration in Mussel Tissue 84 



XI 



LIST OF FIGURES 

FIGURE TITLE PAGE 

1 Clarks Cove Station Locations 6 

2 Station Description 7 

3 Clarks Cove Combined Sewer Overflows and Storm Drain 11 
Locations 

4 Clarks Cove Temperature 15 

5 Clarks Cove Dissolved Oxygen 15 

6 Clarks Cove Salinity 15 

7 Mean Mussel Shell Growth by Station and Deployment 19 

8 Mussel Tissue Mercury Concentration 22 

9 Mussel Tissue Zinc Concentration 23 

10 Mussel Tissue Nickel Concentration 24 

11 Mussel Tissue Lead Concentration 25 

12 Mussel Tissue Copper Concentration 26 

13 Mussel Tissue Cadmium Concentration 27 

14 Mussel Tissue Chromium Concentration 28 

15 Mussel Tissue Arsenic Concentration 29 

16 Comparison of PCB Tissue Concentration 33 

17 Lawrence Experiment Station vs. Division of Marine 42 
Fisheries Values as a Percent of EPA "Mega Mussel" 

Values 



Xlll 



INTRODUCTION 



In 1987 the Massachusetts Division of Water Pollution Control (DWPC), Department 
of Environmental Protection (DEP) applied for and received funding from the U.S. 
Environmental Protection Agency (EPA) Buzzards Bay Project to conduct a pilot 
biomonitoring program in Clarks Cove, New Bedford, Massachusetts using caged 
mussels ( Mytilus ejdulis) . This study is one of several being conducted in 
Buzzards Bay for the EPA Buzzards Bay Project over the past two years. These 
research projects are diverse and address water quality issues identified as 
being priority concerns in Buzzards Bay, mainly; bacterial contamination, 
nutrient enrichment, and toxic contaminants in fish and shellfish. Information 
gathered during this study phase will be used by the Buzzards Bay Project staff 
to develop a Comprehensive Conservation and Management Plan (CCMP) for Buzzards 
Bay. 

The CCMP will provide strategies for pollution abatement and prevention 
throughout the watershed of the bay. In addition, the CCMP will include 
recommendations for long-term monitoring to assess the effectiveness of the water 
quality clean-up and management techniques that are employed. 

The goals of this project were primarily to address questions relating to water 
quality monitoring techniques. In general, the "pilot" portion of the study was 
to design and implement a simple biomonitoring technique that could be performed 
by local, state, and/or regional agencies that would enable detection of long- 
term spatial and temporal trends in contaminant concentrations. More 
specifically, the study was to provide information that could be used to assess 
trace element and bacterial contamination in the water column of Clarks Cove, 
an area that receives discharges from as many as nine (9) combined sewer 
overflows from the City of New Bedford and flows from seven (7) storm drains from 
Dartmouth and New Bedford watersheds. In addition, the DWPC saw this as an 
opportunity to expand its water quality monitoring capabilities by examining this 
methodology for use as a tool to assess trace element contamination in sea water. 
The Massachusetts state analytical laboratory, the Lawrence Experiment Station 
(LES), does not have a "Clean bench" facility that is necessary to directly 
measure the trace concentrations of heavy metals and metalloids present in sea 
water. 

Historically, the basic goal of water quality monitoring programs was to collect 
chemical and physical data which was used to characterize the general water 
quality of an area (Perry et. a. 1987). The design of many monitoring programs 
today still reflect this often random data gathering "objective", despite the 
fact that the intent and expectations of monitoring programs have matured. 
Monitoring programs are now relied upon to provide sound information on which 
to base management decisions. According to Segar, et. al. (1987) most marine 
monitoring programs have been inefficient or ineffective in providing specific 
information that can be used by the manager. These researchers recommend the 
use of transplanted bioindicator organisms to monitor temporal changes of bio- 
available contaminants in an area. The test animals, suspended in the water 
column, ingest, filter and/or absorb what is biologically available to them, 
providing a time integrated measure of the abundance of specific bio-available 
contaminants . 



Within approximately the past fifteen years, the use of indicator organisms to 
monitor coastal water quality has become widely accepted. These studies have 
used both transplanted (i.e., caged) or indigenous test animals. The most ideal 
organisms for these types of studies appears to be bivalves. Capuzzo et. al. 
(1987) attribute the use of shellfish for these types of studies, particularly 
in monitoring heavy metals, to their metals bioaccumulation ability, sensitivity 
to metals concentration gradients, and importance to large programs such as the 
National Shellfish Sanitation Program and the Mussel Watch Program. They also 
point out, however, that there is no standard methodology for collecting these 
data sets. 

Farrington et. al. (1987) and Tripp and Farrington (1984) presented the following 
comprehensive list of reasons why bivalves are considered the most useful 
organisms for this approach: 

1. Bivalves are widely distributed geographically. This characteristic 
minimizes the problems inherent in comparing data for markedly different 
species. 

2. They are sedentary and are thus better than mobile species as integrators 
of chemical pollution in a given area. 

3. They have reasonably high tolerances to many types of pollution, in 
comparison to fish and Crustacea. 

2 5 

4. They concentrate many chemicals by factors of 10 to 10 compared to 
seawater in their habitat making trace constituent measurements easier 
to accomplish in their tissues than in seawater. 

5. An assessment of biological availability of chemicals is obtained. 

6. In comparison to fish and Crustacea, bivalves exhibit low or undetectable 
activity of those enzyme systems which metabolize many xenobiotics such 
as aromatic hydrocarbons and PCBs. Thus, a more accurate assessment of 
the magnitude of xenobiotic contamination in the habitat of the bivalves 
can be made. 

7. They have many relatively stable, local populations that are extensive 
enough to be sampled repeatedly, providing data on short and long-term 
temporal changes in the concentrations of pollution chemicals. 

8. They survive under conditions of pollution that often severely reduce or 
eliminate other species. 

9. They can be successfully transplanted and maintained on subtidal moorings 
or on intertidal shore areas where populations normally do not grow - 
thereby allowing expansion of areas to be investigated. 

10. They are commercially valuable seafood species on a worldwide basis. 
Therefore, measurement of chemical contamination is of interest for public 
health considerations. 



Another advantage of using mussels and oysters that is relative to this 
particular study is that these animals can integrate pollutant levels over space 
and time, an advantage over sampling seawater and sediment for pollution 
assessment that can provide only very short-term (via seawater) or long-term (via 
sediments) contaminant integration (Goldberg, 1986). 

Specific advantages of using transplanted animals taken from a relatively 
unpolluted site and suspended in cages in the test area over sampling indigenous 
animals for contaminants are (de Kock and van het Groenewoud, 1985): 1) the 
animals are derived from a common stock, thereby reducing a potential source of 
variability when comparing geographical locations; 2) the period of exposure to 
the environment is known and can be controlled; 3) monitoring locations can be 
chosen, regardless of whether or not the animals occur there naturally. 

The EPA conducted a study in 1982 to evaluate the use of caged mussels to monitor 
ocean disposal of municipal sewage sludge in the New York Bidge (Phelps et. al., 
1982). The study concluded that the use of transplanted caged mussels as a 
biomonitoring tool in coastal waters was feasible. Some of the large scale 
national water quality monitoring programs employing bivalves include the EPA 
Mussel Watch Program, which was conducted at over 100 sites around the coast 
during 1976-1978, and the current National Status and Trends Mussel Watch Program 
being conducted by National Oceanic and Atmospheric Administration (NOAA) on 150 
coastal sites. In the United Kingdom, mussel watch programs were conducted from 
1977-1979 at over two hundred sites along the coastlines of England, Wales, 
Scotland, and Ireland. 

There are also more localized bioaccumulation studies using indicator organisms 
designed to monitor a specific point source. For example, the EPA has required 
bioaccumulation assessment plans to be included in several recent NPDES permits. 
These plans call for the use of Mvtvlis edulis (blue mussel) and Crassostrea 
virginica (eastern oyster) to monitor survivability and contaminant 
bioaccumulation at sites within the zone of initial dilution of the sewage 
outfalls. Massachusetts sewage treatment facilities that are currently 
developing a plan or are already conducting bioaccumulation studies as part of 
their NPDES permit requirement include the Lynn Water and Sewer Commission, 
Swampscott Wastewater Treatment Plant, South Essex Sewerage District (SESD), and 
the Massachusetts Water Resources Authority (MWRA) . The EPA provides a guidance 
document entitled, "Methods for Use of Caged Mussels for In Situ Biomonitoring 
of Marine Sewage Discharges" (1983) that they recommend for use when designing 
bioaccumulation studies for these permits. Also in Massachusetts, caged mussel 
studies conducted by the New England Aquarium (1986, 1988) have been included 
as part of environmental impact studies to aid in the design and siting of ocean 
outfalls for SESD and MWRA. 

It is evident from the literature that this methodology has become widely used 
and accepted by researchers as well as environmental regulators. The Buzzards 
Bay Technical Advisory Committee (TAC) has recognized the importance of this 
technique in the development of a coastal monitoring program that would be 
capable of detecting water quality trends in space and time. The monitoring 
effort in Buzzards Bay requires efficient techniques that will enable scientists 
to characterize long-term temporal and spatial water quality changes that result 
from point and/or nonpoint pollution abatement strategies and/or deleterious 
activities that may occur within the watershed. Although biomonitoring guidance 
documents do exist (U.S. EPA, 1983), there still is no single, widely accepted 



standard operating procedure for conducting these types of bioaccumulation 
studies. More over, there appears to be even less agreement on how to interpret 
the results. With these problems and the needs of DWPC and the Buzzards Bay 
Project in mind, this pilot study was designed to address the following 
objectives: 

1. To evaluate the impact of urban point sources of contamination into 
Buzzards Bay by assessing concentrations of selected trace elements and 
coliform bacteria in the tissues of the blue mussel (M. edulis) that have 
been suspended in cages at three sites located along a transect 
originating in Clarks Cove, New Bedford. 

2. To compare shell growth between stations in a percentage of the test 
animals. 

3. To examine the feasibility of this type of bio-indicator study as a water 
quality monitoring technique for the Division of Water Pollution Control. 

4. To conduct an inter-laboratory calibration exercise with the Division of 
Marine Fisheries to demonstrate the degree of variability between 
laboratories that may be encountered in a study of this kind. 

This report also contains the results of mussel tissue PCB analysis, although 
this task was not included in the biomonitoring study funded by EPA. Results 
are reported and briefly discussed in this report mainly because the task was 
an integral part of this pilot study and the information it provides will be used 
by DWPC to assess the usefulness of this technique for monitoring PCB 
contamination in other coastal areas of Massachusetts. 



METHODS AND DESCRIPTION OF STUDY SITE 



Study Design 

Arrays cages were deployed at three stations oriented along a north-south 
transect originating in Clarks Cove, New Bedford and extending approximately 5.6 
km (3.5 mi) in a south-south easterly direction out into Buzzards Bay (Figure 
1). Station A was located near the head of Clarks Cove. Station B was 
established at the mouth of the cove midway between the eastern and western 
shorelines. Station C was located in Buzzards Bay near Nun #4 LR approximately 
1.7 miles northeast of Round Hill Point in Dartmouth. Water depth at Station 
A and B at low tide was approximately 5 meters and low tide depth at Station C 
was approximately 9 meters. 

By establishing stations in a land to seaward direction a contamination gradient 
was expected to be observed, with highest levels of metals and bacteria predicted 
in tissues collected from Station A at the head of the cove nearest the urban 
sources (i.e., combined sewer overflows and storm drains), and lowest levels 
anticipated in tissues from reference Station C located over 1 mile (1.6 km) 
located away from land based pollution sources. (See pages 10 - 13 for a 
complete description of the study site. ) Before establishing these station 
locations it was important to consider the influence of currents in the study 
area. Although little information has been published on the hydrodynamics of 
Clarks Cove, available research results supported the selection of a north-south 
transect on which to locate stations. In the main body of Buzzards Bay the 
currents are complex. Net displacement of a particle over a tidal cycle is about 
102 km (EG&G for CDM) . Signell (1987) characterized the circulation pattern in 
the bay as tidally dominated. Wind is also an important mechanism determining 
subtidal circulation especially in shallow embayments and estuaries. EG&G's 
survey described tidal currents in the New Bedford Clarks Cove area as running 
generally south to north-northeast into Clarks Cove on the flood tide and north 
to south-southwest on the ebb tide. 

Each station was located in an area of soft bottom sediments indicating that 
deposition, rather than scouring was taking place. This also enabled sediments 
to be collected for analyses from each site and helped maintain similarity 
between stations. The cage assembly was anchored by one or two 8"xl6" cinder 
blocks and attached to floating lobster buoys to mark their location. This 
design was identical to that used by the EPA, Environmental Research Laboratory 
(ERL) in Narragansett, RI for similar caged mussel biomonitoring studies they 
have conducted in New Bedford Harbor (Don Phelps and William Nelson, EPA, ERL, 
Narragansett, RI, personal communication). With this design, field personnel 
were able to set out and retrieve the cages from a boat rather than rely on scuba 
divers to access the cages. Each cage contained twenty-five (25) mussels 
( Mvtilus edulis) with total shell lengths all between 5-7 cm. Figure 2 
illustrates the design of the cage array for one station. For the growth study 
ten of the twenty-five animals in one cage of each replicate were marked with 
an individual number etched in the shell surface (methods employed for the growth 
study are described below). Each station was made up of four replicates. Each 
replicate consisted of 50 animals divided equally into two cages for a total of 
200 animals per station. A typical exposure period, from deployment to 
collection lasted about sixty days with a new group of mussels set out each time. 




\ 



FIGURE 1 



BUZZARDS BAY CAGED MUSSEL 

PILOT BIOMONITORING STUDY Oct.1987-Sept.l988 



Clarks Cove Station Locations 



FIGURE 2 



BUZZARDS BAY CAGED MUSSEL 

PILOT BIOMONITORING STUDY Oct.l987-Sept.l988 

Station Description: 4 Replicates per Station, 
2 Cages per Replicate, 25 Mussels per Cage, 
200 Mussels per Station 




The EPA (1983) recommends a 30 day exposure time for metals bioaccumulation 
studies whereas de Koch and van het Groenewoud (1985) state that some metals may 
require over 150 days to bioaccumulate in mussels. After discussions with the 
Buzzards Bay Technical Advisory Committee and personnel from Woods Hole and EPA, 
ERL, Narragansett the 60 day exposure period was selected. This allowed for 
twice the EPA recommended exposure time. Longer periods were rejected to avoid 
or minimize the degree of fouling that may occur on the cages and to reduce cage 
loss due to wear and tear from extended periods of weathering. The one year 
study period that began in October thus allowed for five, 60 day exposure 
periods, or deployments, that occurred on the following dates: 

First deployment - October 29, 1987 - January 13, 1988 

Second deployment - January 13, 1988 - March 16, 1988 

Third deployment - March 16, 1988 - May 11, 1988 

Fourth deployment - May 11, 1988 - July 13, 1988 

Fifth deployment - July 13, 1988 - September 21, 1988 

Field and Laboratory Procedures 

The same field procedures were followed for each deployment period. Blue mussels 
(M. edulis ) were collected by hand by Division of Water Pollution Control (DWPC) 
personnel at low tide from a tidal creek near the town beach in Sandwich, MA. 
Immediately after collection, a subset of these animals were sent, on ice, to 
the Lawrence Experiment Station (LES) for baseline tissue analysis. These 
baseline samples consisted of the following: four replicates of 15 animals each 
were placed in labeled, sterile plastic bags for trace element tissue analysis 
(As, Cd, Cr, Cu, Hg, Ni, Pb, Zn) . Twenty-five mussels were placed in a labeled 
sterile plastic bag for total and fecal coliform bacteria tissue analyses. 
Although not funded as part of this study, four replicates of 15 animals each 
were wrapped in aluminum foil and labeled for PCB and PAH analysis. The samples 
for the organics analysis were taken to the DWPC laboratory in Westborough and 
frozen for later analysis at the LES. The remaining mussels were transported 
in clean, plastic-lined coolers to the EPA Environmental Research Laboratory in 
Narragansett, RI. At this lab the animals were placed in flow-through seawater 
tables and left overnight. The following morning the mussels were sorted by size 
and 120 animals in the 5-6 cm range were selected for the growth study. These 
mussels were consecutively numbered from 1 to 120 using a dremel drill to etch 
the surface of the shell. The longest portion of the shell was measured to the 
nearest 0.1 mm using a Manostat (model 5921) caliper. The same individual 
performed all of the shell measurements with same caliper throughout the study. 
This technique was similar to that followed by personnel at the EPA, ERL, 
Narragansett, RI (William Nelson, EPA, Narragansett, RI, personal communication). 

Twenty-five mussels were placed in each cage, which was appropriately labeled 
by station and replicate. One cage per replicate contained 10 numbered animals 
among the twenty-five. Lids were secured with small plastic tie wraps. Cages 
were secured to the trawler float with heavy duty tie wraps for easy removal. 
The cages were left in the flow through seawater tables overnight. 

The following morning the mussel cages were transported in coolers to Clarks 
Cove, New Bedford. All stations were accessed by boat. At each station the 
cages from the previous deployment were retrieved and the new replicates were 
deployed. The replicates were spaced approximately 25 meters apart. 



8 



At each station water samples were collected with a van Dorn grab sampler 1 meter 
below the surface and 1 meter from the bottom. 

These samples were collected to assess whether basic environmental conditions 
were similar at each site, as well as to make sure these conditions were suitable 
for mussel survival. Water samples to be analyzed for total solids, suspended 
solids, chlorides, and turbidity were collected in clean polypropylene 
containers. Samples to be analyzed for total phosphorus, orthophosphate, 
ammonia, and total Kjeldahl-nitrogen were collected in clean, acid rinsed bottles 
and acidified to pH 2.0 with 2 ml of 50 percent H 2 S0 4 . Samples for chlorophyll 
a analysis were collected in clean polypropylene containers. All samples were 
tagged for identification and stored on ice in coolers for transport to the LES 
laboratory. 

Temperature, dissolved oxygen, salinity, and conductivity in the water column 
at each station were measured with a Hydrolab Surveyor II. Data and field 
observations pertaining to weather, sea conditions, and test animal and cage 
conditions after retrieval were recorded in a bound field notebook. On several 
occasions a YS1 Model #33 SCT meter was used to measure temperature, conductivity 
and salinity; dissolved oxygen was measured according to the Winkler technique. 
(Refer to Appendix A for details of meter accuracy, and sample treatment 
methods. ) 

On September 21, 1988 one sediment grab was collected at each station with a 
petite ponar grab dredge (Karlisco International Corp., El Cajon, Ca 92002). 
Prior to sampling the dredge was rinsed in seawater to remove any residual 
sediment. The inside of the dredge was then rinsed with reagent grade acetone, 
followed by a rinse with reagent grade hexane, followed by a final rinse with 
seawater. All chemical rinse waste was collected and transported back to the 
laboratory for proper disposal. The dredge contents were emptied into a plastic 
tray and subsamples of the sediment were scooped into separate specially cleaned 
16 oz. glass, screw-top, wide-mouth jars prepared for metals and organics. Care 
was taken to prevent the collection of sediments in direct contact with the tray 
and/or sides of the dredge. All samples were identified with tags and stored 
on ice in coolers for delivery to LES. See Appendix A for details of sample 
bottle preparation. 

The sediments were analyzed at the LES for the following parameters: Trace 
elements (as total metals or metalloids): As, Cd, Cr, Cu, Hg, Ni, Pb, Zn; 
percent total volatile solids; PCBs and PAHs. 

Appendix A presents the methodology employed at the LES for the analysis of the 
various water and sediment quality parameters. 

After collection the cages were left unopen and placed on ice in coolers for 
transport back to the DWPC laboratory in Westborough. The following day the 
cages were opened and the numbered animals were measured and individual shell 
length was recorded. The number of animals that were dead were noted along with 
the degree of fouling on the cages and on the animals themselves. Dead animals 
were identified by empty shells or by a strong odor of decay. Fifteen mussels 
were randomly selected from each replicate group and were placed in sterile 
plastic bags identified for trace element analysis. Fifteen animals were wrapped 
in aluminum foil and labeled and stored in the freezer (at 4°C) for later PCB and 
PAH analysis, and the remaining mussels (depending on how many were lost due to 



mortality) were placed in labeled sterile plastic bags for total and fecal 
coliform bacteria analysis. The samples for trace element and bacteria analysis 
were then immediately transported on ice to the LES. 

Methods of tissue preparation and analysis for trace element and bacteria in 
shellfish employed at the LES are outlined in Appendix A. 

Inter-Laboratory Calibration 

An inter-laboratory calibration exercise was carried out between the 
Massachusetts Division of Marine Fisheries (DMF) and the Lawrence Experiment 
Station. The DMF proposed to analyze mussel tissue homogenate samples for trace 
elements: As, Cd, Cr, Cu, Hg, Ni, Pb, Zn at their Cat Cove Marine Laboratory 
in Salem, MA. A portion of the same tissue homogenate prepared by the LES for 
trace element analysis was frozen and stored at the laboratory for later delivery 
by DWPC personnel to the DMF. In September of 1988 the DMF notified DWPC that 
it would tissue homogenate samples from LES. 

Ten samples were delivered to DMF on October 3, 1988. Appendix D contains a 
complete description of the DMF project plan and analytical procedures followed 
at the Cat Cove DMF laboratory. 

On March 28, 1989 the LES and DMF were each given 3 replicate frozen samples of 
a standard mussel tissue homogenate ("mega mussel") prepared by the EPA. Both 
laboratories were requested to analyze the tissue homogenate for the same suite 
of eight heavy metals and metalloids using the same methodologies employed during 
the caged mussel study. 

Description of Study Site 

2 2 
Clarks Cove is small, with a surface area of 5.18 km (2 mi ) and an average 

water depth of 5 meters at MLW. The drainage area for the cove is comparatively 

large with the majority (approximately 8.1 km or 2,000 acres) lying within the 

City of New Bedford. The remaining watershed (approximately 2.4 km or 500 

acres) is located within the boundaries of the Town of Dartmouth. Almost 94 

percent of the total New Bedford drainage area is served by combined sewers (CDM, 

1983). 

Along the shoreline of the cove there are nine combined sewer overflow (CSO) 
outfalls and seven storm drain pipes (Figure 3). Table 1 lists each CSO and 
its location and description. 

CDM estimated that 961 million gallons of storm and untreated wastewater were 
discharged to Clarks Cove in 1983. Forty-three percent (or 413 million gallons) 
of this was from CSO discharges, 6 percent (58 million gallons) was from dry 
weather discharges and 51 percent was from storm water runoff. They estimated 
that CSO discharges occur on an average of 75 times a year and they come from 
two major active outlets at the head of the cove (CSO #003 and #004). 



10 



FIGURE 3 



BUZZARDS BAY CAGED MUSSEL 

PILOT BIOMONITORING STUDY Oct.1987-Sept.1988 



Clarks Cove Combined Sewer Overflow Outfalls ICDM,1983) 
and Storm Drain Locations 



<?#? 




TABLE 1 
CLARKS COVE COMBINED SEWER OVERFLOWS 1 



CSO OUTLET DIAMETER 

NUMBER LOCATION (Inches) 

003* Cove Road and Padanaram Ave. 54" 

004 Hurrican Barrier Pumping Station 96" x 84" 

005* Dudley Street and West Rodney 18" 

French Blvd. (W.R.F. Blvd.) 

006 Lucas Street and W.R.F. Blvd. 24" 

007 Capitol Street and W.R.F. Blvd. 24" 

008 Calamet Street and W.R.F. Blvd. 18" 

009 Aquidneck St. and W.R.F. Blvd. 18" 

010 Bellevue St. and W.R.F. Blvd. 12" 
0101 Hudson St. and W.R.F. Blvd. 18" 



* Contaminated by dry weather sanitary flow from storm drains 
connected to the outfall, as observed by CDM (1983). 

CDM Interim Summary Report on CSO Phase I, December 1983 



12 



The dry weather discharges occur as a result of structural or maintenance related 
problems of the existing sewer system. For example, the dry weather flow at CSO 
#004, estimated at over 0.16 MGD, is caused by a plugged dry weather connection. 
Historically, the highest coliform densities in Clarks Cove have been in the 
northern sector of the cove, presumably because of CSO dry weather discharges. 
The waterbody is classified as SA in accordance with the Massachusetts Water 
Quality Standards, but these standards are violated frequently. Clarks Cove 
receives heavy recreational use in the form of swimming, fishing, and boating. 
There are two public beaches and one private beach, and several boat ramps 
located around the cove. The cove is closed to commercial fishing and 
shellf ishing. Beach closings are reportedly rare. 



13 



RESULTS 



Water Quality 

The physical and chemical water quality data collected during the study year are 
presented by station and date in Appendix B. Figures 4-6 illustrate the seasonal 
trend of temperature, dissolved oxygen, and salinity measured at one meter above 
the bottom at the three station locations. As shown, these parameters fluctuated 
similarly at each station throughout the survey year. 

Salinity at all stations ranged between 27 - 32.2 parts per thousand during the 
year. Dissolved oxygen values ranged from a low of 5.0 mg/1 measured at Station 
A in July to high of 12.8 mg/1 measured at Station C in March. The July 
dissolved oxygen values exhibited the greatest between station differences (5.0 
mg/1 at Station A and 7.2 mg/1 at Station C). 

Temperature, salinity and dissolved oxygen concentrations were within ranges 
necessary for mussel growth and survival at all of the stations. 

Nutrient concentrations measured at the stations during the study were low to 
moderate and fell within ranges reported in the Buzzards Bay water quality 
surveys (MDWPC, 1985, 1986a), with the exception of Station B during March. This 
station exhibited elevated suspended solids and turbidity as well as high total 
Kjeldahl-nitrogen, total phosphorus and orthophosphate concentrations in the 
bottom water column sample. It is possible that the sediments were disturbed 
during sampling and this contaminated the sample. Suspended solids and turbidity 
were otherwise low and within expected ranges. These parameters followed similar 
trends between stations throughout the year. 

Sediment Quality 

Table 2 presents the sediment trace element, PCB, PAH, and percent total volatile 
solids data for each station. All sediment samples were collected on September 
21, 1988. A rigorous assessment of the sediment quality was beyond the scope 
of this study. Since the results cannot be normalized, and only one sediment 
grab per station was collected, an in-depth comparison and evaluation of sediment 
quality cannot be made from these data. 

Station A sediments contained the highest concentrations of all trace elements, 
and organics measured, with the exception of nickel, which was slightly higher 
at Station C (6.5 mg/km versus 5.5 mg/km at Station A). Zinc and PCB 1254 
concentrations were above category III dredge spoils criteria (MDWPC, 1983) at 
Station A. Arsenic was also elevated at this site. Station B and C sediments 
contained similar concentrations of most of the trace elements, and results were 
within ranges reported in the Buzzards Bay sediment survey (MDWPC, 1985-86). 
PCB 1254 concentration was higher at Station B (exceeded Category III criteria) 
than at Station C (Category II). 

PAH concentrations were relatively low at all of the stations, but the greatest 
number of compounds (and concentrations) were found at Station A and the least 
at Station C. 

Percent total volatile solids were similar at all stations. 



14 



Oct 
1987 



FIGURE 4 

CLARKS COVE TEMPERATURE 

Stations A, B, and C 



Jan 
1988 



March May 

Sampling Month 




Sept 



O 14 
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Oct 
1987 



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CLARKS COVE DISSOLVED OXYGEN 

Stations A, B, and C 



Jan 
1988 



March May 

Sampling Month 



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Stations A, B, and C 



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15 



TABLE 2 

CLARKS COVE 

SEDIMENT DATA 

TRACE ELEMENTS, POLYCHLORINATED BIPHENYLS, POLYCYCLIC AROMATIC HYDROCARBONS 

AND PERCENT TOTAL VOLATILE SOLIDS 1 

September 21, 1988 



PARAMETER 



STATIONS 
B 



CATAGORY III 
DREDGE SPOILS 
CRITERIA 2 



Trace Elements (mg/kg dry weight) 



Arsenic 

Cadmium 

Chromium 

Copper 

Mercury 

Nickel 

Lead 

Zinc 

PCB 1254 (pg/g) 

Percent Total Volatile Solids 



2.4 


1.4 


2.0 


>20 


6.5 


<1.0 


1.0 


>10 


41 


23 


29 


>300 


60 


60 


24 


>400 


0.335 


0.170 


0.105 


>1.5 


5.5 


2.5 


6.5 


>100 


90 


33 


25 


>200 


500 


85 


90 


>400 


2.3 


1.3 


0.91 


>1.0 


5.9 


5.4 


6.7 


_ 



PAH (pg/g dry weight) 



Benzo ( a ) anthracene 

Benzo ( a ) pyrene 

Bnezo(k) f luoranthene 

Chrysene 

Fluoranthene 

Phenanthrene 

Pyrene 



0.80 


0.20 


- 


0.57 


- 


- 


0.96 


- 


- 


0.56 


0.13 


- 


1.10 


0.41 


0.20 


0.55 


0.18 


0.10 


1.10 


0.32 


0.20 



Total PAHs reported by LES 



5.64 



1.24 



0.50 



See Appendix A, Table A-5 for methods of analysis and limits of detection. 



DWPC, 1983 



16 



Cage Loss and Mussel Mortality 

Percent mortality that occurred at each station during each deployment is 
presented in Table 3. The number of cages (replicates) lost during each exposure 
period is also listed in this table. The percent mortality was calculated by 
dividing the total number of dead animals found at a station by 200 (the total 
number of animals deployed at each station) and multiplying by 100. Mortality 
was usually very low, generally only 0-4 animals per station were lost. However, 
during the last exposure period of 7/13-9/21 mortality was very high (25-53 
percent). An extreme degree of fouling by barnacles and algae was observed on 
the cages and animals themselves from this period. Also, several small starfish 
were found in many of the cages. Cages collected from all other deployments 
exhibited very little fouling and no starfish were observed inside them. 

Four cages were lost during two of the deployment periods. Other periods 
experienced only a loss of 1 or 2 cages. One replicate (C4) lost during the 
first deployment period was recovered on 9/21/88 at the same site after almost 
one year of exposure. Out of the original 50 animals, 27 survived from this 
group. 

Shell Growth 

Mean shell growth and standard deviation for animals at each station and for each 
deployment are shown in Figure 7. The average shell growth over a 60-day period 
of 120 mussels is highly variable as illustrated by the standard deviation bars 
(one S.D.) on the graph. This variability masks any statistical differences that 
may exist between Stations A, B, and C for any one deployment period. However, 
from the graph it appears that mean shell growth at these stations exhibit fairly 
similar trends during each period. The largest differences in shell growth are 
seen between exposure periods, although these are not statistically significant 
due to the large standard deviations. As expected, in general, the spring and 
early summer exposure groups show the largest increase in average shell growth, 
and the fall and winter periods produce the least amount of growth. 

Tissue Bacteria Concentrations 

Tissue total and fecal coliform bacteria concentrations are presented in Table 
4. Tissue samples from the last deployment period were not analyzed for bacteria 
concentration due to high mortality resulting in an insufficient number of live 
mussels available for the analysis. It was felt that the bacteria analysis was 
the most expendable of the parameters, because tissue bacteria data obtained from 
the last four deployments were erratic and did not supply any more useful 
information for monitoring long-term trends in bacteria contamination than could 
be obtained from direct water column sampling techniques (see discussion 
section) . 

Baseline tissue bacteria concentrations were generally much higher than tissue 
concentrations measured in animals after exposure, indicating that the Sandwich, 
MA site may not be appropriate for collecting "clean" mussels if bacterial 
contamination is a concern. A large number of birds were often observed near 
the area where the mussels were collected. 



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In addition, bacteria concentrations between stations for each deployment did 
not exhibit a discernable pattern. It was expected that animals nearest the head 
of the cove would accumulate the highest bacteria concentrations. This was not 
the case. On several occasions, Station C, the reference site located out in 
Buzzards Bay, had the highest bacteria counts. In general, if total coliform 
was high (>1,000 colonies per 100 ml), fecal coliform was also elevated. 

Tissue Trace Element Concentrations 

Figures 8 through 15 illustrate the results from the tissue analysis for trace 
elements. Concentration is reported in mg/kg (wet weight) for mercury (Hg) , 
chromium (Cr) , cadmium (Cd), arsenic (As), lead (Pb), nickel (Ni), copper (Cu), 
and zinc (Zn) . 

Each graph illustrates the tissue concentration of one trace element over all 
of the deployment periods. The bars represent the mean tissue concentration of 
the metalloid of all the replicates for each station, grouped by deployment 
period. One standard deviation is depicted on the graph to illustrate the 
variability of the data about the mean. Appendix B contains the tissue trace 
element concentration data as reported by the LES. Results from each deployment 
were examined separately. Comparison of contaminant concentration throughout 
the year is not possible since a new set of animals was used for each 60 day 
deployment period. Inter-exposure period comparisons of this nature would only 
have been possible if all of the animals had been deployed at the beginning of 
the study and subsampled throughout the year. 

Statistical analysis using the nonparametric Kruskal-Wallis test (Zar, 1984) 
was performed on the tissue trace element concentration data. Nonparametric 
statistics were chosen because the variances of the groups of data being compared 
were not homogeneous. Under these conditions this nonparametric ANOVA test is 
more powerful than the one-way ANOVA (Zar, 1984). The Kruskal-Wallis statistic 
tested the null hypothesis that trace element concentration in tissue from the 
baseline station and Stations A, B, and C were the same. (H Q : [metalloid] is 
the same at all stations.) 

Appendix C contains sample statistical calculations. Table 5 presents a summary 
of the results of the nonparametric ANOVA tests. 

A significant difference between mean trace element concentration was detected 
at the 95% confidence level for only 13 of the 35 groups of data tested. (During 
deployments four and five, detection limits of Cd, Cr and Pb were increased as 
a result of a change in laboratory procedure. As a consequence almost all values 
were reported as less than detection limits for these periods, thus limiting 
further analysis and comparison of these data sets.) 

Since the Kruskal-Wallis multiple comparison test does not indicate where the 
significant differences occur in the data set, a nonparametric Tukey-type 
multiple comparison test was applied to locate where the differences existed 
(Zar, 1984) for these 13 data sets. (See Appendix C for sample calculations.) 
Table 6 summarizes the results of these calculations. 

Due to high standard error values in several of the data sets only 8 of the 13 
Tukey tests detected significant differences between the means. 



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29 



TABLE 5 



SUMMARY OF KRUSKAL-WALLIS NONPARAMETRIC ANALYSIS OF VARIANCE 



TRACE 
ELEMENT 



DEPLOYMENT DEPLOYMENT DEPLOYMENT DEPLOYMENT DEPLOYMENT 
1 2 3 4 5 



Zinc 



accept H reject H accept H reject H reject H 



Mercury accept H Q accept H Q accept H Q accept H Q accept H c 



Nickel 



accept H Q accept H Q reject H Q accept H Q accept H Q 



Cadmium accept H 



Chromium accept H 



Arsenic 



reject H 



accept H Q 
accept H Q 
reject H Q 



accept H Q 
accept H Q 
reject H 



can not 
analyze 

can not 
analyze 



can not 
analyze 

can not 
analyze 



reject H reject H 



Lead 



Copper 



reject H accept H reject H 



can not 
analyze 



Hypothesis being tested: 



accept H 



reject H accept H Q accept H Q reject H Q accept H Q 



H Q : The mean trace element concentration of baseline = Station A = 
Station B = Station C 



30 



TABLE 6 



SUMMARY OF TUKEY-TYPE NONPARAMETRIC MULTIPLE COMPARISON TEST 



DATA SET 



RESULTS 



Zinc deployment 2: 
Zinc deployment 4: 
Zinc deployment 5: 



Baseline different (lower) than Sta. A,B,C; but A,B,C same 
Baseline different (lower) than Sta. B; but all others same 
Baseline different (higher) than Sta. C; but all others same 



Nickel deployment 3: No significant differences detectable due to large standard 

error 



Arsenic deployment 1: No significant differences detectable due to large standard 

error 
Arsenic deployment 2: Baseline different (lower) than Sta. C, but all others same 
Arsenic deployment 3: Baseline different (lower) than Sta. B, but all others same 
Arsenic deployment 4: Baseline different (lower) than Sta. A, but all others same 
Arsenic deployment 5: Baseline different (higher) than Sta. A, but all others same 



Lead deployment 1: No significant differences detectable due to large standard 

error 
Lead deployment 3: Baseline different (higher) than Sta. C, but all others same 



Copper deployment 1: No significant differences detectable due to large standard 

error 
Copper deployment 4: No significant differences detectable due to large standard 

error 



31 



In every case, the significant differences in the means were due to the baseline 
mean tissue trace element concentration being different from one or more of the 
other stations (A, B, or C) . Usually, but not always, baseline concentrations 
in these cases were lower. 

In four out of five of the exposure periods arsenic baseline tissue 
concentrations were significantly different from either Station A, B, or C. In 
three of these data sets arsenic was lowest in the baseline samples. However, 
since neither Station A, B, or C were consistently highest (or lowest) throughout 
the study, spatial patterns of arsenic distribution in this area are not evident. 

Baseline concentration of zinc for three out of five exposure periods was 
significantly different from Station A, B, or C. However, as with arsenic, 
consistent spatial patterns of distribution of zinc at these stations cannot be 
detected nor can further speculation as to what may be causing these differences 
be made. 

For lead, the baseline concentration was significantly higher than Station C for 
one exposure period. 

Tissue concentration of cadmium, chromium, mercury, nickel and copper were not 
significantly different for any of the exposure periods. 

None of the test Stations (A, B, or C) exhibited significant differences in trace 
element tissue concentration indicating differences in bioaccumulation of these 
elements were not spatially significant for these stations. This suggests that 
trace element concentration available for uptake in the water column at these 
stations was not significantly different between Station A, B, or C. 

PCB Tissue Concentrations 

The results of the PCB analysis of tissue from deployments 1, 3, 4, and 5 are 
illustrated in Figure 16. Mean values of the data normalized with percent lipids 
are shown on the graph. Appendix B lists the PCB tissue concentrations as 
reported by LES. Only arochlor 1254 was detected in any of the tissue samples. 
Percent lipid concentration for each sample is also reported in Appendix B. 

The lowest PCB concentrations were consistently measured in the baseline mussel 
tissue and the highest PCB concentrations were found in tissue from Station A. 
The next highest PCB concentrations were found at Station B and relatively low 
concentrations of PCB were usually detected in tissue from Station C. 

Interlaboratorv Calibration Exercise 

The results from the interlaboratory calibration exercise between the Department 
of Environmental Protection's laboratory (LES) and the Division of Marine 
Fisher ie's laboratory (DMF) at Cat Cove, Salem, MA are presented in Table 7. 



32 



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3A 



Values for cadmium and chromium are not comparable because the detection limits 
of the LES analyses were much higher than the DMF's detection limits. The DMF 
reported "not quantifiable" concentrations of mercury with the values falling 
between 0.006 mg/kg and 0.020 mg/kg. This range is less than or near the 
detection limit reported by the LES for mercury analysis. 

For copper, nickel, zinc and lead the values reported by the LES were 
approximately five times higher than that reported by the DMF for the same tissue 
homogenate samples. 

On March 28, 1989 standard mussel tissue samples prepared by the EPA laboratory 
in Narragansett, RI were hand delivered to the Lawrence Experiment Station and 
the Division of Marine Fisheries laboratory at Cat Cove, Salem, MA. Results of 
each laboratory's analyses are presented in Table 8. 



35 



TABLE 8 

INTERLABORATORY CALIBRATION RESULTS 

U.S. EPA STANDARD "MEGA MUSSEL" TISSUE (mg/kg dry weight 1 ) 

METAL 



AGENCY 



Cd 



Cr 



Cu 



Ni 



Pb 



Zn 



U.S. EPA, Narragansett, RI 

(Average tissue metals concentration) 



2.08 



2.15 



12.8 



6.84 



9.11 



135 



Lawrence Experiment Station 



1.9 



1.9 



11.7 



6.2 



8.3 



119 



U.S. EPA range of values' 



1.99- 1.91- 12.2- 6.37- 7.94- 126- 
2.18 2.36 13.8 7.24 10.25 142 



Division of Marine Fisheries 



2.17 



1.98 



11.7 



8.06 



90 



LES obtained dry weight of sample by drying homogenate for 2 days at 90 °C 
and weighing entire sample. 

J LES results are reported as an average of 2AA analyses (except for Pb - 
only enough sample for one analysis). 

3 

U.S. EPA analyzed 50 samples to obtain range and average tissue metals 
concentration of the standard homogenate. 

Division of Marine Fisheries results were converted to dry weight by 
multiplying wet weight values by 6.83 (EPA's reported wet/dry weight 
ratio for the "mega mussel" homogenate). 



36 



DISCUSSION 



Study Design 

Studies that involve comparisons of selected variables over space and time 
ideally require that all environmental conditions that may affect test results 
be similar either through controlled laboratory conditions or, in field studies, 
as a function of study design. However, too much control placed on the 
experimental design may create an artificial situation which may obscure 
interpretation of the relationship of the data to actual field conditions. For 
this study it was important that the stations selected exhibit very similar 
measurable environmental conditions. The three stations chosen were oriented 
on a north-south transect from the head of shallow Clarks Cove to open water in 
Buzzards Bay; consequently depth was not the same at each location. (5 meters 
at Station A and B vs 9 meters at Station C. ) Despite depth differences however, 
temperature, dissolved oxygen and salinity were essentially the same at each 
station supporting the assumption that all of the animals were most likely 
exposed to similar environmental conditions during each deployment period. 

Growth (as measured by average shell length increases over the exposure period) 
and mortality were not significantly different between stations which also 
indicates that the environmental conditions necessary for mussel growth and 
survivorship at each site were the same. 

If growth differences between sites were evident in this study, then differences 
in tissue trace element bioaccumulation between sites (if present) would be more 
difficult to interpret and could not necessarily be attributed solely to 
available contaminant concentrations in the water column. 

Enseco, Inc. (1990) reported that mussels deployed near sewage treatment outfalls 
in Boston Harbor that survived appeared to be generally healthier than reference 
site organisms. Based on these findings, assumptions that more polluted sites 
would negatively affect the health (and growth) of test animals cannot be made. 

Although not performed in this study other methods of growth or condition 
assessment may be more effective than simple shell length measurements. A 
practical method of determining a body condition index should be investigated 
and, if at all possible, applied in future caged mussel studies of this kind. 

Mortality was usually very low except for the last exposure period where 
predation by starfish was suspected to have caused the 25-53 percent mortality 
observed in the cages. Although it is not known if starfish predation on 
bivalves occurs seasonally in Buzzards Bay it may be wise to avoid deploying the 
mussels in cages during this time of year in this particular area. For all but 
the last deployment, the low mussel mortality assured sufficient numbers of 
animals for tissue analysis. In addition, similar mussel growth, mortality and 
environmental conditions found at each station reduces sources of variation that 
may influence spatial differences in contaminant uptake by the mussels. 



37 



Coliform Contamination 

The use of caged mussels to monitor coliform contamination over time and space 
was ineffective. Since the animals clear their gut in approximately 24-48 hours, 
any assumptions regarding temporal changes in bacteria concentration in the water 
column are limited to one day time periods. Furthermore, the potential for 
encountering variability within the stations is high due to the fact that the 
animals are filter feeders, and may each be filtering different volumes of water 
over a 24-hour periods and thus ingesting highly variable amounts of bacteria 
over this relatively short time. For this reason, monitoring of coliform 
bacteria to detect long-term changes in water column bacteria densities should 
not be performed via tissue concentrations. 

Monitoring whole mussel tissue bacteria concentration is potentially a valid 
technique for making spatial comparisons of bacteria concentrations in the water 
column at discrete time periods. However, the method is much more labor 
intensive, results are highly variable, and it offers only slight advantage (i.e, 
from a temporally non-integrated grab sample of water versus a 24-48 hour time 
integrated tissue sample) over simple, direct water column bacteria sampling 
methods. 

Trace Element Concentration 

As is evident from the data, tissue trace element concentration was extremely 
variable, not only statistically between replicates at each station, but 
spatially and temporally as well. Due to variation within the data set 
significant differences in trace element concentrations, if they existed in the 
water column at any of these stations over time, were not usually detectable. 
The magnitude of trace element bioaccumulation in the mussel tissue was small 
in comparison to this variability. It is important to examine the major factors 
that may influence the variability of the data and its resulting usefulness. 

The often large variances of the station replicates as well as the differences 
in average tissue trace element concentration between Stations A, B and C 
(spatial differences) and between baseline mussel tissue and Stations A, B, and 
C (temporal differences) may be the result of any one, or a combination of the 
following factors: 1) natural seasonal variability; 2) data bias or errors 
resulting from field study design and implementation; 3) data bias or errors 
resulting from laboratory procedures; 4) actual temporal or spatial differences 
in water column trace element concentrations. 

Natural seasonal variation can account for as much as 15-60 percent of the 
variability in observed values (Capuzzo, et. al., 1987). Seasonal variation may 
be a result of the physiological state of the animals, environmental conditions, 
and metal speciation and bioavilability (Capuzzo, et. al., 1987). Seasonal 
variability would not influence the between station (spatial) differences of the 
data because comparisons of these results were made between mussel tissue from 
the same exposure period. As previously discussed, results indicated that these 
mussels were experiencing similar seasonal environmental and physiological 
conditions as measured by temperature, dissolved oxygen, salinity, chlorophyll 
a concentrations, and shell growth at each site. 

Tissue trace element concentrations were not compared at each station over 
several exposure periods. With this study design, comparisons of this type would 



38 



be weak because discrete groups of animals were set out and measured each 
exposure time, rather than subsampled periodically from a large group that had 
been exposed for the entire study year. However, seasonal variability may have 
caused differences between baseline tissue concentrations and Stations A, B, and 
C since baseline animals were collected in Sandwich at the beginning of the 
exposure period approximately 60 days earlier than the animals they were compared 
to from Clarks Cove. 

Percent lipids were not measured in the tissues homogenized for trace element 
analyses; except for growth, no other parameters were measured to assess the 
physiological condition of the mussels. Percent lipids were measured in tissue 
homogenate prepared for organics analysis (see Appendix B) . Although not 
assessed during this study, spawning condition of the animals is known to be 
directly related to whole animal percent lipid concentration. Lipid 
concentration increases as animals prepare to spawn and drops sharply after 
spawning. Spawning reportedly leads to loss in tissue weight, increase in 
percent water and decline in condition indices. Prior to spawning lipid-rich 
gametes may contain higher concentrations of lipophilic organic contaminants and 
lower concentrations of heavy metals than somatic tissues. After spawning a drop 
in organic concentrations and an increase in metal concentrations may result 
(Robinson and Ryan, 1988). Therefore, to greatly enhance tissue data 
interpretation future caged mussel studies should include an assessment of the 
spawning condition of the animals. This should be made at the time of 
deployment, when baseline trace element tissue concentrations are measured, and 
when the animals are retrieved after the exposure period. Inferences about 
adverse impacts of toxic trace elements on the health of the mussel cannot be 
made, although this factor may have also been responsible for some variability 
of the data. Animals showed an average increase in shell length for each 
exposure period. Average shell increases were the largest during the third and 
fourth deployments (March 16 - May 11 and May 11 - June 16, respectively) . No 
correlation between growth and trace element concentrations can be made. The 
goals of this study did not include an attempt to relate contamination 
concentration with indications of stress in the organism. 

As previously discussed, the field study design attempted to equalize as many 
between station environmental variables as possible. The study design may 
benefit from including at least one more replicate at each station since the 
variances between the four replicates were often high. In addition, it has been 
suggested that not all of the animals receive equal exposure time bunched-up in 
the square cages. Flat cages that spread the animals into one-layer would allow 
all to have more of a chance to filter equal amounts of water. Cages of this 
design were not available for this study. To reduce the likelihood of this type 
of bias animals were selected randomly from the bunches in the cages when 
preparing the sample bags for the laboratory. Other studies performed with 
square cages did not report evidence of this type of bias (Robinson and Ryan, 
1986, 1988 and Nelson, personal communication). 

Besides the systematic or random variability introduced via seasonality and field 
study design, data variability introduced through laboratory procedures must be 
considered an important factor when interpreting the results. The Lawrence 
Experiment Station analyzes samples in "batches." QA/QC tests are performed on 



39 



a percentage of samples from each batch. The QA/QC results during this study 
were acceptable, suggesting that variation between stations and/or replicate 
samples was due to other factors (i.e., the effects of seasonality, or 
differences in contaminant concentrations). 

Determination of dry weight concentrations of the trace element was not requested 
as part of this study. However, water content is extremely variable in these 
animals, not only seasonally but individually, and will definitely affect the 
calculation of the results. Ideally, dry weight should be determined separately 
for each sample homogenate prepared, rather than using an average dry weight of 
mussel tissue to normalize the data. Robinson and Ryan (1988) state that in 
transplant studies it is impossible to determine whether metal body burdens 
actually changed as a result of exposure if changes in tissue weights were not 
monitored. They report that changes in mussel tissue weight can be assessed by 
measurements of tissue dry weight, condition index and gonadal index. Future 
tissue biomonitoring studies should include a determination of tissue dry weight 
to reduce data variability. 

Possible sources of data variability were discussed with LES personnel and they 
included procedures in sample preparation and analytical methodologies. Some 
of these sources can be minimized with the use of a more efficient method of 
tissue homogenation and/or via procedural modifications such as determining the 
dry weight of samples and using consistent sample sizes for analysis throughout 
the study. 

Results of tissue metals concentration from this study and ranges of values 
reported for several other metals bioaccumulation studies are compared in Table 
9. Arsenic concentration was not measured in the other studies listed here, so 
it is not included in this comparison. Mercury, cadmium, and chromium 
concentrations fall within the ranges reported by other researchers. Mercury 
concentration never approached the US Food and Drug Administration limit of 
1 /jg/g wet weight. Cadmium and chromium were also very low, often below the 
detection limits of the analyses, and concentrations never fluctuated much from 
site to site, nor did they vary over exposure times. From this study, it appears 
that mercury, cadmium and chromium either require a longer exposure period to 
bioaccumulate in the mussel or there were low concentrations of bioavailable 
metal in the water column at these sites, de Kock and van het Groenewoud (1985) 
report that cadmium accumulation is a slow process requiring about 150 days to 
reach equilibrium values. These researchers were also unable to demonstrate 
differences in mercury concentration from several sites in 60 day transplant 
studies. Robinson and Ryan (1988) state that transplanting clean mussels to 
polluted sites to assess seawater contaminant levels is only successful when 
metals concentrations are high enough to result in appreciable bioaccumulation. 

Maximum concentrations of lead, copper, nickel, and zinc greatly exceeded ranges 
reported from other studies (see Table 9). Station A tissue most often contained 
the highest metals concentrations; however as previously discussed, between 
station differences of tissue concentrations of these metals could not be 
detected due to large within station variances. In general, seasonal peaks in 
Cu, Ni, Pb and Zn tissue concentration occurred more in the late spring and early 
summer (also the period when the greatest shell length increases were measured). 

Possible reasons for these extremely high values of Zn, Cu, Ni, and Pb includes 
laboratory sources of variability discussed previously as well as the natural 



40 



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41 



FIGURE 17 

Plot of Lawrence Experiment Station (LES) v* 

D.v.s.on of Marine Fisheries (DMR Values 

as a Percent of EPA "Mega-Mussei" Va, ues 

1 10 



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42 






variability of actual metals concentration and varying rates of bioac cumulation 
and regulation of each metal by Mvtilus edulis during different times of the 
year. 

No range of arsenic tissue concentrations were available for comparison. Ranges 
of concentration for this element were not included in Table 9 for this reason. 
For arsenic the within station variances were usually lower than for other 
metals. Arsenic concentrations from July and September samples were 
significantly higher at all stations than at any other time of year. 

Interlaboratorv Calibration Exercise 

The interlaboratory calibration exercise with the DMF yielded differences in 
tissue wet weight concentrations of copper, nickel, and zinc from aliquot s of 
the same sample homogenate. Values of cadmium, chromium, and lead reported by 
DMF were lower than the detection limits established by LES in their analysis 
therefore these metals data were not comparable. DMF obtained unquantif iable 
concentrations of mercury (0.006 ppm<x<0.02ppm) which were also not comparable 
to the LES values. 

The LES values of Cu, Ni, and Zn were on the order of 5-6 times higher than the 
DMF results. Discussions with the LES and DMF concluded that differences in 
their results could not be definitively explained. It is difficult to attribute 
the differences in results to the analytical equipment because samples were 
exposed to variations in handling before being extracted for analysis. It was 
not the goal of this exercise to isolate and test for variability in analytical 
equipment, otherwise complete sample preparation would have been performed at 
only one laboratory. Galloway, et. al. (1983) found that identical techniques 
in different laboratories do not necessarily give similar results. They report 
that homogenates of several matrices prepared by two different agencies and 
subjected to intercomparison exercises have consistently shown wide ranging 
results. For comparison of methodologies used by each laboratory please refer 
to Appendices A and D. 

In an attempt to assess whether these differences were actually due to 
differences in laboratory techniques and not in the samples themselves, each 
laboratory was requested to analyze standard EPA mega mussel tissue homogenate. 
Results show concentrations obtained by the LES are within 9-12 percent of the 
values reported by EPA (Figure 17). It is interesting to note that the LES 
values are all slightly less than EPA average concentrations. The LES results 
for Pb and Cr fall within the range of values reported by EPA. Cd, Cu, Ni, and 
Zn values were only slightly less than the minimum EPA values. DMF results for 
Cd, Cr, and Pb fall within EPA's reported ranges. As illustrated in Figure 17, 
values for Cd, Cr, Cu, and Pb reported by DMF are within 4-11 percent of EPA 
averages. DMF did not report results for nickel. Zinc concentration obtained 
by DMF was comparatively low however, differing by more than 30 percent from the 
EPA average. 

Considering that the EPA averages were based on a sample size of 50 and the LES 
and DMF values were derived from an average of two (or less) analysis, results 
for these trace element analyses appear to be in very good agreement among the 
laboratories. Both laboratories tended to have a bias toward lower values as 
compared to EPA results but the reasons for this trend are unknown. 



43 



Based on the results of the "mega mussel" interlaboratory calibration exercise, 
no evidence for why the results of the study mussel tissue interlaboratory 
analyses were so different between LES and DMF can be found. 

Tissue Concentrations of PCBs and PAHs 

This project only required that a portion of the animals from each exposure 
period be archived (frozen) for future organics analysis. However, since the 
organics laboratory at the LES was able to perform the analysis on many of the 
archived samples during the study period, the results are presented and briefly 
discussed as part of this report. 

No PAHs were detected in the tissue samples from either Clarks Cove or Sandwich, 
MA. In contrast, Capuzzo, et. al. (1987) report mussel tissue collected from 
a variety of sites in New England, including Cape Cod, contained detectable 
levels of PAHs. Eisler (1987) however, found in general that PAHs show little 
tendency to biomagnify in food chains. He attributed this to the fact that most 
PAHs are rapidly metabolized. Specific reasons for PAHs not being detected in 
this study cannot be offered. An interlaboratory comparison between the LES and 
EPA, Narragansett or Woods Hole Oceanography Institute organics laboratories may 
provide some insight as to what is happening here. 

PCB tissue concentrations were normalized by the percent lipid concentration of 
the sample to account for differences in PCB concentration created by differences 
in lipid content of the tissue. As seen from Figure 16 the results show a 
consistent pattern of higher PCB concentrations in the tissues from Station A 
to decreasing amounts in Station B and even lesser amounts in Station C. Not 
only are spatial differences evident, but differences can be seen between 
baseline and test site concentrations for each deployment period. From this 
consistent pattern it appears that 60 day exposure periods allow sufficient time 
for bioaccumulation of measurable amounts of PCBs in mussels. EPA recommends 
at least 30 days (U.S. EPA, 1983), although differences in PCB concentration of 
test animals have been detected after just 2 weeks of exposure in New Bedford 
Harbor (W. Nelson, personnel communication) . 

Based on the well documented PCB contamination in New Bedford Harbor it is not 
surprising that PCB concentrations at Station B as well as Station A, were 
relatively high. The area that encompasses both stations has been closed to 
bottom fishing and lobstering by the Department of Public Health due to PCB 
contamination. None of the tissues from this study contained PCBs in excess of 
the FDA action level of 2.0 \iqlq. Concentrations ranged from <0.04-1.2 p/g/g. 
This range falls within that observed in US Mussel Watch data from Cape Cod and 
Buzzards Bay. In New Bedford Harbor the range of PCB tissue concentration from 
Mussel Watch data was much higher (3.08-6.86 pg/g) (Capuzzo, et. al., 1987). 

In this study the use of Mytilus edulis as a sentinel organism to monitor PCB 
contamination in the water column appears more successful than for monitoring 
metals contamination. The standard deviations of the station replicates were 
low and concentration averages followed an expected pattern for every deployment. 
Sediment PCBs also followed a similar, relative concentration gradient. Results 
appear to be more straightforward to interpret both spatially and temporally. 
Also by normalizing with percent lipids much of the variability that may have 
been introduced as a result of differences in reproductive condition of the 
animals was eliminated. 



44 



SUMMARY 

The use of caged mussels for coastal biomonitoring proved to be a very feasible 
field technique from the standpoint of available resources at the Technical 
Services Branch of DWPC. Questions that remain should be addressed through 
increased communication with the analytical laboratory, continued interlaboratory 
calibration exercises, and modification of the study design. Based on the 
results and suggestions from other researchers, several modifications of the 
study design and analytical procedure are recommended: 1) trace elements that 
exhibited low bioconcentration should be eliminated from the study (Cd, Cr, and 
Hg); 2) tissue dry weight should be determined for each sample homogenate; 3) 
the sample should be thoroughly homogenized; 4) interlaboratory calibrations 
should continue with sample tissues from the study sites as well as with a 
standard tissue homogenate (EPA mega mussel); 5) increase focus on using this 
technique to monitor PCB contamination; 6) examine the effect of longer exposure 
periods by subsampling from a large group of transplanted mussels over a one year 
period; and 7) the method should not be used to monitor coliform bacteria 
contamination . 

In most of Buzzards Bay, metals contamination is most likely not high enough to 
bioaccumulate to statistically significant amounts. If definitive 
bioaccumulation was not measured at Clarks Cove, other less impacted areas would 
be less expected to show significant bioaccumulation of tissue in trace element 
concentrations. From this study it is evident that actual differences, either 
spatial or temporal would have to be very large to be significant. However, 
this study as well as others indicate that temporal and spatial characterization 
of changes in PCB contamination are possible using caged mussels. Serious 
consideration should be given to using this technique as part of a LONG-TERM 
monitoring program in Buzzards Bay, especially in the New Bedford area. 

It is important that biomonitoring studies such as this continue to be developed 
and performed by agencies responsible for water quality monitoring. Of the three 
basic methods used to assess pollutants in the coastal environment; water 
sampling, sediment sampling, and sampling of biota, the later has received the 
least attention by the Massachusetts Division of Water Pollution Control. The 
bioavailability of contaminants however, should be a major concern, not only 
because it can provide a means of determining time-integrated pollutant 
concentrations but because of the long-term implications to human health, and 
more important, the overall health of the ecosystem. Although water pollution 
standards today are based on measurements of water and sediment, a contaminant 
can only be considered a threat to the environment if it can be taken up by the 
biota. 



45 



BIBLIOGRAPHY 



American Public Health Association 1985 <u- an ** ^ „ ^ 

Q { »at er „, ,..^.>„. _ ..i^n ys; f :^; ^ 

Camp Dresser and McKee. 1983. city of New Bedford t - • 

combined Sewer Overflow. Phase I. TJ*J>° ' I " t " 1 " SU ™ ar * R **° rt ° n 



Capuzzo, J.M., A. McElroy, and G. Wallace 1987 »- v, 

218 » Street. ..,^£££H£ "^Bpp. ** """"" R ^°"< 

Filling in ^^^^^x^SST'^rS! """^ "s^* 1 ' "" 

Commonwealth of Massachusetts, Division of Water Pn ,i <- ■ 

-«*. Bay Mater Quality survey Da - j^^s^rs- oxsix 85 - 

y ourvey uata, Part A. Westborough, MA 01581 

^^^^e^r^:^ 1 ^ 011 - W " er — " — -as. 

0X58X. 9 Procedures - Biomomtoring Program. Westborough, MA 

de Kock, w. Chr. and H. van het Groenewoud loft* v, ,, ,,. 

and Elimination Dynamics of s™» Z t'- -." Modelling Bioaocumulation 

Based on "in Situ-^e"atfonsTi t h „^ P ° Uutants < cd ' ■». FCB, HCB, 
,or society, Heport ^^ XOS^if^^f ,' ^X 1 ""^ 

Report . 85(1.11). 81pp. " eView * U ' S ' Fish Wildlife Ser v ice Rim 

June 1990. Marblehead, MA 10945 M ° nit ° rin 9 *«*"». "89 Deployment. 

Farrington, J.W., A.C. Davis, B.W. Trior, D y »>,., 

"Mussel watch - Measurement, of Cb^aXpo^tanL f" p* 11 ^ 
indicator of Coastal Environmental Quality » He „ a v Bivalves As One 

Aouati. ssaaa ASTM tal ^l^', "*"_; teBtaasiisa to Mnnii-n.-i ,,,, 

Testing and Materials, Phila., ppl25-X 3 V ' ftmerican s ° ci <*y «« 

Farrington, j.w. and G.c. Medeiros ISA* * , 

Analysis for Petroleum Hvdt^V k Evaluation of Some Methods of 
Proceedings f £ ^£ erence On^reve'nl ""^ ° r9anisms - Gained in 
coastal Research Center woods! Lie hT C °° tr ° 1 ° f ° U Poll ^°»- 



46 



BIBLIOGRAPHY (CONTINUED) 



Galloway, W.B., J.L. Lake, D.K. Phelps, P.F. Rogerson, V.T. Bowen, J.W. 
Farrington, E.D. Goldberg, J.L. Laseter, G.C. Lawler, J.H. Martin, and R.W. 
Risebrough. 1983. The Mussel Watch: Intercomparison of Trace Level 
Constituent Determinations. Environmental Toxicology and Chemistry . Vol. 
2. pp395-410. 

Goldberg, E.D. 1986. The Mussel Watch Concept. Environmental Monitoring and 
Assessment . Vol. 7 (1986) 91-103. 

Perry, J. A., D.J. Schaeffer and E.E. Herricks. 1987. "Innovative Designs for 
Water Quality Monitoring: Are We Asking the Questions Before the Data Are 
Collected?", New Approaches to Monitoring Aguatic Ecosystems . ASTM STP 940 
T.P. Boyle, Ed. American Society for Testing and Materials. Phila. pp28- 
39. 

Phelps, D.K., W.B. Galloway, B.H. Reynolds, W.G. Nelson, G. Hoffman, J. Lake, 
C. Barsycz, F.P. Thurberg, J. Graikowski and K. Jenkins. 1982. Evaluation 
Report: Use of Caged Mussel Transplants for Monitoring Fate and Effects 
of Ocean Disposal in the New York Bight Apex. US EPA Environmental Research 
Laboratory, Narragansett, ERIN No. 586. 35pp. 

Robinson W.E. and D.K. Ryan. 1986. Bioaccumulation of Metals, Polychlorinated 
Biphenyls, Polyaromatic Hydrocarbons and Chlorinated Pesticides in the 
Mussel, Mvtilus edulis L., Transplanted to Salem Sound, Massachusetts. A 
final report submitted to Camp Dresser and McKee, Inc. 20 October 1986. 

Robinson W.E. and D.K. Ryan. 1988. Bioaccumulation of Metal and Organic 
Contaminants in the Mussel, Mytilus edulis . Transplanted to Boston Harbor, 
Massachusetts. In Project Report submitted to Camp Dresser and McKee, Inc. , 
by the Edgarton Research Laboratory, New England Aquarium, 15 February 1988. 
205pp. 

Segar, D.A., D.J.H. Phillips., and E. Stamman. 1987. "Strategies for Long-Term 
Pollution Monitoring of the Coastal Oceans," New Approaches to Monitoring 
Aguatic Ecosystems . ASTM STP 940, T.P. Boyle, Ed. American Society for 
Testing and Materials, Phila. ppl2-27. 

Signell, R.P. 1987. Tide and Wind-Forced Currents in Buzzards Bay, MA . Woods 
Hole Oceanographic Institute, Woods Hole, MA WHOI-87-15. 86pp. 

Tripp, B.W. and J.W. Farrington. 1984. Using Sentinel Organisms to Monitor 
Chemical Changes in the Coastal Zone: Progress or Paralysis. Proceedings 
of the Ninth Annual Conference of the Coastal Society. Oct. 14-17, Atlantic 
City, NJ. 

U.S. Environmental Protection Agency. 1983. Methods for Chemical Analysis of 
Water and Wastes. EPA-600/4-79-020. 

U.S. Environmental Protection Agency. 1983. Methods for Use of Caged Mussels 
for in situ Biomonitoring of Marine Sewage Discharges. EPA-600/4-83-000. 



47 



BIBLIOGRAPHY (CONTINUED) 



U.S. Food and Drug Administration. 1988. Food and Drug Procedure. Pesticide 
Analytical Manual. January. Washington, D.C. 

Zar, J.H. 1984. Biostatistical Analysis , 2nd Edition. Prentice-Hall Inc., 
Englewood Cliffs, NJ 01632. 718pp. 



48 



APPENDIX A 



FIELD METHODOLOGY 

AND 

LAWRENCE EXPERIMENT STATION LABORATORY METHODOLOGY 



49 



TABLE A-l 



COMMON SAMPLE TREATMENT METHODS 



PARAMETER 



SAMPLE VOLUME 



SAMPLE 
CONTAINER 



1 



IMMEDIATE SHIPBOARD 
PROCESSING & STORAGE 



Dissolved Oxygen 
Temperature 

Specific Conductance 

Total Solids 

Suspended Solids 

Chloride 

Total Kjeldahl-Nitrogen 

Ammonia-Nitrogen 

Total Phosphorus 

Orthophosphate 

Turbidity 

Chlorophyll a/ 
Phytoplankton 



300 ml (2) 



1 1 (2) 

1 1 (2) 

1 1 (2) 

1 1 (2) 

500 ml (2) 

500 ml (2) 

500 ml (2) 

500 ml (2) 

1 1 (2) 
200 ml 



G (1) 
" (1) 

P/G (1) 

P/G (1) 
P/G (1) 
P/G (1) 
G (1) 
G (1) 
G (1) 
G (1) 
G (1) 
P/G (1) 



MnS0 4 ; KI: no sunlight/ 
or (4) "in situ." 

In situ recorded to 
nearest 0.1°C/F or (3), 
(4), (5) 

"In situ" reading/or cool 
4°C (3), (4) 

Cool 4°C 

Cool 4°C 

Cool 4°C 

H 2 S0 4/ pH <2.0, cool 4°C 

H 2 S0 4 , pH <2.0, cool 4°C 

H 2 S0 4 , pH <2.0, cool 4°C 

H 2 S0 4/ pH <2.0, cool 4°C 

Cool 4°C 

Cool 4°C (5) 



G - Glass 
P/G - Polypropylene or glass 

(1) Required containers, preservation techniques, and holding time, per Table 
II 40 CFR Part 136. 

(2) Massachusetts Division of Water Pollution Control, Technical Services Branch, 
Engineering Section, Standard Operating Procedures. 

(3) Yellow Springs Instrument, Model 33-S-C-T meter and probe. Yellow Springs 
Instrument Co., Inc., Yellow Springs, Ohio 45387. 

(4) Hydrolab Surveyor II, Model SVR2-SU sonde unit, Model SVR2-DV Digital read 
out. Hydrolab Corp., P.O. Box 50116, Austin TX 78763. 

(5) Massachusetts Division of Water Pollution Control, Technical Services Branch, 
Biomonitoring Program 1988, Standard Operating Procedures. 



50 



TABLE A-2 
PARAMETER AND COLLECTION METHODS EMPLOYED AT SEDIMENT STATIONS 



PARAMETER 



SAMPLE VOLUME 

(Liters) SAMPLE CONTAINER 



IMMEDIATE FIELD 
PROCESSING & STORAGE 



PCB 1016/1242 Sediment 2(25-100 g) 



G/Aluminum Foil 
Septum 



Cool to 4°C 



PCB 1248 Sediment 



2(25-100 g) 



G/Aluminum Foil 
Septum 



Cool to 4°C 



PCB 1254 Sediment 



2(25-100 g) 



G/Aluminum Foil 
Septum 



Cool to 4°C 



PCB 1260 Sediment 



2(25-100 g) 



G/Aluminum Foil 
Septum 



Cool to 4°C 



PAHs Sediment 



2(25-100 g) 



G/Aluminum Foil 
Septum 



Cool to 4°C 



Metals Sediment 



25-100 g 



G/Teflon Septum 
or Plastic Wrap 
Septum 



Cool to 4°C 



G = Glass 



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58 



TABLE A- 7 
METHOD FOR CHLOROPHYLL a ANALYSIS (MDWPC, 1988) 

3.7.1 DEFINITION ; Chlorophyll is a pigment found in plants that allows the 
organism to use radiant energy for converting carbon dioxide into organic 
compounds in a process called photosynthesis. Several types of 
chlorophylls exist and these and other pigments are used to characterize 
algae. One type, chlorophyll a, is measured for it is found in all 
algae. A knowledge of chlorophyll a concentrations provides qualitative 
and quantitative estimations of phytoplanktonic and periphytic biomasses 
for comparative assessments of geographical, spatial and temporal 
variations. 

3.7.2 EQUIPMENT NEEDS 

1. Fluorometer - either Turner 111 or the Turner Design 10-005-R field 
fluorometer is used. They must be equipped with blue lamp F4T5. 

Corning filter -5-60-excitation 
Corning filter - 2-64-emission 
Photomultiplier 

2. Tissue grinder and tube - Thomas Tissue Grinder 

3. Side arm vacuum flask and pump 

4. Millipore filter holder 

5. Glass fiber filter: Reeve angel, grade 934H, 2.1 cm 

6. Centrifuge (Fisher Scientific Safety Centrifuge) 

7. 15 ml graduated conical end centrifuge tubes with rubber stoppers 

8. 90% aqueous acetone 

9. IN HCL 

10. Saturated magnesium solution in distilled water 

11. Test tube racks 

12. Borosilicate cuvettes - Turner 111 - 3" cuvettes 

Turner Design - 8" cuvettes 

13. Aluminum foil 

14. Test tube brushes - conical end 

15. Parafilm 



59 



TABLE A-7 (CONTINUED) 



3.7.3 LOG- IN PROCEDURE 

As samples are received they are logged in and assigned a number. The 
samples can be frozen for further analysis, or the filter ground up for 
analysis the following day. 

3.7.4 SAMPLE PREPARATION 

Samples are generally processed as soon as they come into the laboratory, 
unless there are extenuating circumstances, such as faulty equipment 
and/or time constraints. Samples not to be analyzed within 24 hours are 
frozen for future analysis. 

The procedure for freezing samples follows: 

1. Label a 2-inch Whatman petri dish with the sample number using an 
indelible pen. 

2. Using tweezers, take a 2.1 cm Reeve Angel, grade 934AH, glass fiber 
filter and place it on the Millipore filtering flask screen. Do 
not touch the filter. Attach the glass tube to the filter flask 
with the metal clamp. 

3. Shake the sample well. 

4. Measure out 50 mis of sample or less. If an amount other than 50 
mis is used it should be recorded in the chlorophyll data book. 

5. Pour the measured sample into the filter tube and turn on the 
vacuum. The sample should pass quickly through the glass fiber 
filter; therefore more of the sample should be added. If the sample 
is not filtering through - either because too much sediment is 
present or the algal concentration is too high - then less than 50 
mis can be filtered. A notation is made in the chlorophyll data 
book which lists the amount that was filtered. 

6. Unclamp the filter holder and with tweezers transfer the filter to 
the previously marked petri dish. 

7. Cover the petri dish and wrap it in aluminum foil to keep out the 
light. The petri dish with the glass fiber filter is then stored 
in the freezer. 

8. Return the sample bottle to the refrigerator if algal counts or 
identifications are requested. 

9. Rinse the graduated cylinder and filter holder in distilled water. 



60 



TABLE A-7 (CONTINUED) 

3.7.5 ANALYTICAL PROCEDURE 

1. Follow steps 2-6 under "Sample Preparation." 

2. Filter 50 ml (or less if necessary) of sample through a glass fiber 
filter under vacuum. 

3. Push the filter to the bottom of tissue grinding tube. 

4. Add about 3 ml of 90% acetone and 0.2 ml of the MgC0 3 solution. 

5. Grind contents for 3 minutes. 

6. The contents of the grinding tube are carefully washed into a 15 
ml graduated centrifuge tube. 

7. Bring the sample volume to 10 ml with 90% acetone. 

8. Test tubes are wrapped with aluminum foil and stored in the 
refrigerator for 24 hours. 

9. Test tubes are taken out of the refrigerator and put into the 
centrifuge. 

10. Test tubes are then centrifuged for 20 minutes and the supernatant 
decanted immediately into stoppered test tubes. 

11. Tubes are allowed to come to room temperature. The temperature is 
recorded and the samples are poured into a cuvette (3" for Turner 
111 and 8" for Turner Design) . 

12. The Turner 111 requires a warm-up period of at least one-half hour, 
while the Turner Design 10-005-R does not require a warm-up period. 

13. With Turner 111, use a blank of 90% aqueous solution of acetone to 
zero the instrument. Open the front door of the fluorometer and 
put in the cuvette containing the 90% acetone and close the door. 
Press the start switch. The dial should move back to 0; adjustments 
can be made with the calibration knob. This process should be 
repeated as often as necessary, i.e., if the blank is not staying 
on zero; but no alteration should be made until a series of samples 
is completed. 

14. The Turner Design must also be zeroed to an acetone blank. The 
sample holder is located at the top of the Turner Design field 
fluorometer and should be recovered with the black cap after the 
sample is put in it. 



61 



TABLE A- 7 (CONTINUED) 



15. Readings for both the Turner 111 and the Turner Design should be 
within 20-80% of the scale. This can be achieved by either reducing 
or increasing the opening to the lamp by moving the knob on the 
right front of the Turner 111 fluorometer. The sensitivity levels 
are lx, 3x, lOx, and 30x. The sensitivity level must be recorded 
in the chlorophyll data book in addition to whether the high 
intensity or regular door was used. After the first reading, 2 
drops of 2N HC1 is added to the cuvette. A piece of parafilm is 
used to cover the cuvette which is then inverted four times to mix 
the sample thoroughly. The sample is re-read and the new value 
recorded. 

16. The procedure for the Turner Design field fluorometer is basically 
the same as for the Turner 111. The sample is put into the cuvette 
holder and the manual switch used to go from one sensitivity level 
to the next without opening the door. A reading of between 20-80% 
is still required for accuracy. Readings are taken before and after 
acid is added to the sample. The level of sensitivity (lx, 3x, 6x, 
lOx, 31. 6x) must be recorded in the chlorophyll data book, as well 
as whether the levels were set at 1 or 100. 

Calculation of Chlorophyll Concentrations 

Chlorophyll concentrations are determined by using the following 
formulas: 

chlorophyll (fig/1) = Fs rs (Rb-RA) 

rs-1 



pheophytin (/ug/l) = Fs rs (rsRa-Rb) 

rs-1 

where, 

Fs = conversion factor for sensitivity level "s" 

rs = before and after acidification ratio of sensitivity level "s" 

Rb = fluorometer reading before acidification 

Ra = fluorometer reading after acidification 

A computer program is used to calculate the chlorophyll concentrations 
for samples run on the Turner Design fluorometer. This program requires 
the investigator to type in the sensitivity level and the difference 
between the before and after acidification values. 

During the summer of 1986 personnel of the Technical Services Branch 
(TSB) conducted a laboratory experiment with a Turner Design Fluorometer 
in order to determine the effect of pheophytin b on freshwater 
chlorophyll a readings. Pheophytin b is the degradation product of 
chlorophyll b which is the primary pigment of green algae. The Turner 



62 






TABLE A- 7 (CONTINUED) 



Design instrument measures the fluorescence of chlorophyll a as well as 
that of pheophytin a and b. Chlorophyll b is not read at the same 
frequency as chlorophyll a. The emission filter used at the TSB (Corning 
C/S 2-64) partially rejects pheophytin b (See: "References" - Turner 
Designs, 1981). It was found and recorded in various unpublished 
memoranda (See "References") that unless a sample had elevated counts 
of green algae the readings obtained prior to acidification and 90 
seconds thereafter would give a reliable estimate of the concentration 
of chlorophyll a in an algal sample. In cases with elevated counts of 
green algae an annotation should be made alongside the chlorophyll a 
concentration stating that the concentration may reflect the presence 
of chlorophyll b and is probably lower than as recorded. As a result 
of this investigation, the TSB now present chlorophyll data as 
chlorophyll a in mg/m . 

3.7.6 INSTRUMENT CALIBRATION 

Fluorometers are calibrated using chlorophyll samples provided by the 
United States Environmental Protection Agency. Calibrations are 
performed at the start of every field season and redone if any changes 
are made to the fluorometer such as changing the light bulb. 

Samples for chlorophyll analysis are periodically split with another 
laboratory or run on two separate fluorometers. 



63 



WET TISSUE DIGESTION FOR METALS ANALYSIS 

BY ATOMIC ABSORPTION SPECTROSCOPY 

AND/OR ICP EMMISSION SPECTROSCOPY 

(FISH, CLAMS, MUSSELS, ETC.) 



CHEMISTRY LAB SOP 
UPDATED 04/13/88 



65 



TABLE OF CONTENTS 

I . Sample Storage 1 

II. Sample Transport 1 

III. Sample Receipt and Recording 1 

IV. Preparation of Glassware 1 

V. Sample Preparation 1 

VI. Sample Weighing 2 

VII. Digestion Procedures 2 

VIII. Sample Digest Filtration 3 

IX. Q.C. Samples 4 

X. Safety Precautions 4 

XI. Glassware, Chemicals, Equipment and Supplies 4 



67 



I. Sample Storage 



a. Fish samples should be wrapped with plastic wrap and stored in 
sealed plastic bags. 

b. Clam and Mussel samples should be stored in sealed plastic bags. 



II . Sample Transport 

a. Samples collected and brought to LES the same day should be 
transported in a cooler with ice. 

b. Samples collected and stored for future delivery to LES should 
be placed in freezer. Samples should be removed from freezer and 
placed in a cooler with ice for delivery. 

Ill . Sample Receipt and Recording 

a. Samples received by the LES Chemical Lab personnel are immediately 
numbered on I.D. Tags and recorded into the Chemistry Lab Log 
Book. 

b. Chain of Custody Samples must be accompanied with approved forms. 

c. Samples are stored in freezer until they are readied for 
processing. 

IV. Preparation of Glassware 

a. All glassware is washed in micronox cleaning solution, rinsed with 
tap water, acid washed with 40% nitric acid solution, and rinsed 
2x or 3x with deionized - distilled water. 

V. Sample Preparation 

a. Remove samples from freezer and thaw. 

b. (1) Fish - The total fish fillet is diced into small sections 

on a nalgene cutting board using a stainless steel knife. 
Transfer the diced fish sections into a 40 oz. or small size 
glass blender top (depending on the amount of sample.) 

(2) Shellfish - Scrub outside of shellfish with a stiff nylon 
bristled hand brush while rinsing under tap water. Shuck 
total clam or mussel sample collected into glass blender top. 

c. Using a variable speed blender start homogenizing sample on low 
speed for 1 or 2 minute intervals (shut off blender between 
intervals to prevent overheating or burning out the blender 
motor) . 



68 



d. Once blender blades start making a uniform contact with sample, 
use higher speeds for 1 or 2 minute intervals. Continue this 
procedure until sample is thoroughly homogenized. 

e. With a teflon spatula transfer homogenized sample to plastic or 
glass container and seal. Label and number. Place in 
refrigerator or freeze samples for up to 6 months. 

Note ; For some large fillets, it may be necessary to split sample 
into aliquots, homogenize separately, and recombined in a clean 
plastic container. Transfer to multi purpose plastic containers, 
label and number. 

f . Rinse knife with deionized - distilled water and wipe clean with 
paper towels. Rinse cutting board with tap water, wash with 5% 
nitric acid solution, rinse 2x or 3x with deionized - distilled 
water, and wipe dry. Clean inside of glass blender and rotor 
blades with hard bristled nylon brush and hot tap water. Rinse 
with deionized - distilled water 2x or 3x. This cleaning 
procedure must be repeated after every sample. 

VI . Sample Weighing 

a. Label and tare 400 ml beaker on balance. 

b. Weigh 10.0 gms of homogenized sample into beaker. 
Note ; Teflon spatula used to transfer sample. 

c. Cover beaker with watch glass. 

d. Record weight to nearest 0.1 gm into Digestion workbook. 

e. For every 10 samples or less a duplicate and spiked sample is 
weighed out. 

Note ; The sample is spiked before digestion using Eppendorf 
pipets and stock 1000 ppm certified standards. Spiked 
concentrations are determined for each batch of samples. 

VII. Digestion Procedure 

a. Add 10 ml concentrated nitric acid to the beaker with sample. 

Note ; Acid should be added under fume hood, safety glasses and 
gloves must be worn. 

b. Cover with watch glass. 

c. Place on steam bath and reflux for 2 hours. 

d. Remove watch glass and evaporate to near dryness. 



69 



e. Add 10 ml concentrated nitric acid and 10 ml of 30% hydrogen 
peroxide (H 2 C 2 ) to beaker. 

f. Cover beaker with watch glass and reflux on steam bath for 2 
hours. 

g. Remove watch glass and evaporate to near dryness. 

h. Add approximately 50 ml of 1% vol/vol hot nitric acid to beaker 
and let stand for 15-30 minutes on steam bath. 

VIII. Sample Digest Filtration 

a. Set up nessler tube (100 ml graduated) in rack with filter funnel 
and #42 Whatman filter paper (18.5 cm). 

b. Wash down filter paper with deionized - distilled water. Discard 
washing from nessler tube. Rinse nessler tube with deionized - 
distilled water. Replace tube in rack with washed filter paper 
and funnel. 

c. Remove beaker from steam bath. While decanting sample into 
funnel, wash sidewalls (inside) and bottom of beakers with 
deionized - distilled water (use a 500 ml side arm wash bottle). 

d. Rinse beaker with two 10 ml aliquots of hot 1% nitric acid 
solution, and filter. 

e. Rinse filter with deionized - distilled water. 

f. Q.S. to 100 ml with deionized - distilled water. 

g. Transfer digest to labeled sample container (125 ml rectangular 
H.D. polypropylene bottle). 

- To ensure thorough mixing, pour digest back into nessler tube, 
and transfer back into sample bottle. 

Note : High density polypropylene sample containers may become 
porous. Acid washing or acid soaking in some cases doesn't remove 
100% of the contaminates adsorbed within the container. 
Therefore, it is recommended that once samples have been 
quantitated, reports have been checked and mailed, that the sample 
containers be discarded. 

IX. Q.C. Samples 

a. A reference standard, duplicate and spiked samples are processed 
through the digestion and filtration procedure for each set of 
10 samples or less. One reagent blank is processed through the 
digestion and filtration procedure for every set of samples. 



70 



X. Safety Precautions 

a. Lab safety practices must be strictly followed. 

b. Protective glasses, gloves, and lab coats must be worn. 

c. Fume hoods should be used whenever necessary. 

d. Safety respirator with acid vapor removal cartridge should be 
worn. 

XI. Glassware, Chemicals, Equipment and Supplies 

a. Glassware 

1. 400 ml beakers (heavy duty) 

2. 50 ml graduated cylinders 

3. Watch glasses 

4. 100 ml graduated nessler tubes 

b. Chemicals 

1. Nitric acid 

2. 1000 mg/L Standards for Atomic Absorption spectrophotometers 
(certified ACS grade) 

3. 30% Hydrogen Peroxide (certified ACS grade) 

c. Equipment 

1. Nalgene filter funnels (10 cm diameter) 

2. Teflon spatulas 

3. 125 ml H.D. Polypropylene sample bottles 

4. 4 oz. and 16 ox. polypropylene multipurpose containers with 
lids. 

5. Repeater pipettors, or automatic dilutors. (500 ml base, 
10 ml delivery) 

6. 500 ml side arm wash bottle 

7. Shellfish shucking knife 

8. Stainless steel fillet knife 

9. Nessler tube rack 

10. Stiff bristled nylon brush (wooden handle) 

11. Nalgene cutting board 

12. Safety glasses 

13. Safety gloves 

14. Safety respirator with acid vapor removal cartridges 

15. Mettler PE1600 Balance 

16. Waring Blender #7012, Model 34BL97, 7 speed 

17. Eberback 40 oz. glass blender with handle (#8442) 

18. Eberback small size glass blender (#8470) 



71 



d. Supplies 

1. Micronox cleaning solution 

2. Plastic Bags (sealable) 

3. Label tape 

4. China marker 

5. Lab coat 

6. Paper towels 



72 



APPENDIX B 
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82 



TABLE B-4 

MUSSEL TISSUE POLYCHLORINATED BIPHENYLS 
(fjg/g wet weight) 

Aroclors 1254 and 1242 



DEPLOYMENT- 




l 


STATION 




REPLICATE 


BASELINE 


A 


B 


C 


1-1 


0.49 


0.95 


0.82 


0.39 


1-2 


ND** 


- 


- 


- 


1-3 


0.15 


- 


- 


- 


1-4 


- 


- 


- 


0.68* 


~x 


0.066 


- 


- 


- 


s 


0.077 


- 


- 


- 


3-1 


0.047 


1.1 


0.40 


0.74 


3-2 


<0.040* 


1.0 


1.1 


- 


3-3 


<0.040 


0.81 


1.0 


- 


3-4 


- 


1.2 


- 


0.62 


"x 


0.423 


1.028 


0.83 


0.68 


s 


0.004 


0.166 


0.38 


0.09 


4-1 


0.058 


0.58 


— 


0.28 


4-2 


<0.040 


0.78 


0.81 


- 


4-3 


0.040 


0.58 


0.68 


0.29 


4-4 


ND 


0.71 


0.62 


0.20 


~x 


0.035 


0.66 


0.70 


0.26 


s 


0.025 


0.10 


0.097 


0.05 


5-1 


0.041 


0.61 


— 


0.45 


5-2 


<0.040 


0.55 


0.94 


- 


5-3 


0.070 


0.65 


0.41 


0.66 


5-4 


<0.040 


- 


0.43 


0.53 


"x 


0.048 


0.60 


0.59 


0.547 


8 


0.02 


0.05 


0.30 


0.110 



- Sample not analyzed 

* Less than values averaged as reported number 
** ND (none detected) values treated as in calculation of 
average. 
*** Tissue exposed from 10/27/87 to 9/21/88. 



83 



TABLE B-5 
PERCENT LIPID CONCENTRATION IN MUSSEL TISSUE 



DEPLOYMENT- 




STATION 






REPLICATE 


BASELINE 


A 


B 


C 


1-1 


2.6 


1.8 


2.2 


2.7 


1-2 


1.7 


- 


- 


- 


1-3 


1.9 


- 


- 


- 


1-4 


- 


- 


- 


1.4** 


X 


2.1 


1.8 


2.2 


2.7 


3-1 


1.4 


2.7 


0.87 


3.0 


3-2 


1.4 


3.0 


2.2 


- 


3-3 


2.8 


1.4 


2.4 


- 


3-4 


- 


1.8 


- 


2.5 


X 


1.9 


2.2 


1.8 


2.75 


4-1 


1.9 


1.3 


— 


0.80 


4-2 


1.4 


1.2 


1.8 


- 


4-3 


1.3 


1.4 


1.4 


1.1 


4-4 


1.1 


1.6 


2.7 


0.83 


X 


1.4 


1.38 


2.0 


0.91 


5-1 


1.8 


1.3 


— 


1.5 


5-2 


1.1 


1.2 


1.9 


- 


5-3 


1.0 


1.1 


0.98 


* 


5-4 


1.3 


- 


1.0 


1.6 


~x 


1.3 


1.2 


1.3 


1.6 



- Sample not analyzed 
* Sample lost in analysis 
** Tissue exposed from 10/27/87 - 9/21/88 



84 



APPENDIX C 
SAMPLE STATISTICAL CALCULATIONS 



85 



KRUSKAL - WALLIS TEST FOR ANOVA 
DEPLOYMENT #4 



Arsenic 



H Q : As concentration is the same in all groups 
H A : As concentration is not the same o< =0.05 

Arsenic 

Base A B C 

0.45 (2.5) 2.2 (6.5) 3.2 (12) 2.4 (8) 

0.37 (1) 4.0 (14) 3.0 (11) 

0.45 (4) 4.5 (15) 2.7 (9) 2.1 (5) 

0.45 (2.5) 3.9 (13) 2.8 (10) 2.2 (6.5) 

n x = 4 n 2 = 4 n 3 = 4 n 4 = 3 

R x = 10 Rj = 48.5 R 3 = 42 R 4 = 19.5 

N = 4+4+4+3 = 15 

2L _E;_ 2 -3 (N+l) 





= 


12 


H 


N(N+1) 




= 


12 




15(16) 




— 


12 



s; 






[ 10 2 + 48 .5 2 + 42 2 + 19. 5 2 ]-3(16) 



[ 25 + 588.06 + 441 + 126.75J-48 



240 
= .05 [ 1,180.81 ]-48 

= 59.04-48 
H = 11.04 



number of groups of tied ranks = 2 
^T = ^(V - tj) C = 1- .ST 



3 



= (2 3 - 2) + (2 3 -2) 



N^-N 



12 = 6 + 6 C = 1- 12 



3,360 
C = 0.9964 



H c = _H_ = 11.04 = 11.0799 
C 0.9964 

H Q 0.05,4,4,4,3 = 7.14 

.*« reject H Q because H c > 7.14 



86 



NON-PARAMETRIC MULTIPLE COMPARISONS 



A Nonparametric Kruskal-Wallis test is applied to Deployment 4 Arsenic values and 
the null hypothesis (they are the same) is rejected. To determine where the 
significant differences occur use: Nonparametric Tukey-type multiple 
comparisons: 



T = 12 



SE = 



N (N + 1) 
12 



12(N-1) 



n> 



n 



B 



SE for n =4,4 



15(16} 



12 



12 



12(14) 



f I + I 

,4 4/ 



= /9.9643 = 3.157 



SE for n = 3,4 = 



15(16) 



12 



12 /l + 1' 
12(14) (3 4J 



= /11.624 = 3.41 



Samples ranked by mean ranks (i) 



Baseline 



4 
(C) 



3 
(B) 



2 
(A) 



rank sums (Rj) 
sample sizes (n,) 



10 



19.5 



42 



48.5 



mean ranks R 



2.5 



6.5 



10.5 



12.13 



Q = R 



% " R A 

SE 



Comparison 



2 vs 1 



Difference 



SE 



12.13-2.5=9.63 



-2o.05,4- 



Conclusion 



3.157 3.050 2.639 Reject 1^: [As] 

different in A 
& Baseline 



2 vs 4 



12.13-6.5=5.63 



3.41 



1.651 2.639 Accept H Q : [As] 

same in A & C 



2 vs 3 

3 vs 1 



do not test 



10.5-2.5=8.0 



3.157 2.534 2.639 Accept H Q : [As] 

same in B & baseline 



3 vs 1 

4 vs 1 



do not test 



6.5-2.5=4.0 



3.41 



1.173 2.639 Accept H Q : [As] 

same in Baseline 
& C 



Overall conclusion: 
Arsenic concentration is different between baseline 
and Station A but the same in all other comparisons 



87 



APPENDIX D 



DIVISION OF MARINE FISHERIES 
PROJECT PLAN AND LABORATORY METHODOLOGY 



89 



QUALITY ASSURANCE PROJECT PLAN 



QUALITY CONTROL SECTION OF THE PILOT 
MONITORING PROGRAM. DEPARTMENT OF ENVIRONMENTAL QUALITY 
AND ENGINEERING, DIVISION OF WATER POLLUTION CONTROL 



FF.EFAFED EY 
COMMONWEALTH OF MASSACHUSETTS , 
DEPARTMENT OF FISHERIES. WILDLIFE. ANO 
ENVIRONMENTAL LAW ENFORCEMENT 



FOR 

U.S. ENVIRONMENTAL PROTECTION AGENCY 
REGION 1 
WATER MANAGEMENT DIVISION 

MAY 28, 1987 
(revised August 19, 1987) 



APPROVALS: 





Mr. W. Leigrr Br idges, <£V inci.pal Investigator x^Date 



Dr. Wenay Wiltse. Buzzards Bay Project Monitor Date 



Mr. Charles Porfert, DeDuty Quality Assurance Officer Date 



91 



TABLE OF CONTENTS 

E2Q§ 

Project !^ame 1 

Projec: Reauesrea By i 

Dare of Reauest 1 

Date of Project Initiation 1 

Project Officer__ 1 

Projec: .Monitor 1 

Quality Assurance Officer i 

P r o j ec : Desc r i d t i en 1 

a. Cc active anc Sccoe 1 

E. Data Usace ' 

C. Design an.G Rationale 1 

D. Mc~ itoring Parameters/Frecuency of Collection 1 

Protect Fiscal Information 2 

Schedule of Tasks ana Procucts 2 

Project Organization and Responsibility 2 

Data Quality Reauirements ana Assessments 3 

Samolm- and Analytical Proceaures 3 

Same I e Custody Procedures 3 

Calibration Proceaures ana Preventive Maintenance 4 

Documentation, Data Reduction, and Reporting 4 

Da t a Va . t ca 1 1 on 4 

Performance ana System Auatts 4 

Correct.ve Action -_ 4 

Reoorts 4 



LIST OF TABLES AND FIGURES 

Table ". . Laboratory Analyses 1 

Table 2. Esttmatea Project Costs 1 

Figure 1. Analysis Recues t Form, Cat Cove 

Marine Laboratory _.___ 6 



CoDtes sent to: Wenav Wilts© (EPA) 

Char les Porfert (EPA) 



W. Leigh Br igges (OMF) 
Jack P. Schwartz (DMF) 
Nina M. Dustcn (DMF) 
Chr is Duerr ing (DEQE) 



92 



1. Project- name: 

Quality Control section for DEQE Pilot Monitoring Program 

2. Project recuested by: 
U.S. EPA. Region 1 

3. Date of reauest : 
Aon I 15. 1987 

4 . Da t e of p r o j ec t i n 1 1 1 a 1 1 on : 
to be determined bv DEGE 

5. Pro lect Of f icer : 

M r . Rona l d Man f r eccn \ a 

6. Pro lect Mon i tor : 
Dr . Wendv W i I tse 

7. Project oescriDticn: 

A. Objective ana scope 

The Division of Water Pollution Contrc: (Deoartment of Envi ronmmental 
Quality and Engineering, Commonweal tn of Massacnuset ts) is conducting 
a monitoring program involving the analysis of the blue mussel, 
Mvt i l is eoul i s, for trace guantities c: arsenic (As), caamium (Cd). 
chromium (Cr), coDDer (Cu). lead (Pb), mercury (Hg)'., nickel (Ni), and 
zinc (Zn) . As Dart of this studv, Cat Cove Marine Laboratorv 
(Division of Marine Fisneries, Department of Fisheries. Wildlife, and 
Environmental Law Enforcement) has the objective of providing auality 
control information on a suPset of mussel samples in order to verify 
trace metal analyses on the larger data Pase and ensure consistency 
Petween sampling periods for the duration of the monitoring program. 



B. Data Usage 
(to Pe determined by DEQE.) 

C. Design and Rationale 
(to be determined bv DEQE) 

D. Monitoring parameters/f reouency of collection 

Mussel swill Pe mom tored for the e i gr : a foremen t i oned metal s 
Samoling will Pe concucted once every two montns ror one year 



93 



E. Parameter Table 



TaDle 1. Laboratory Analyses 



•ameter Matrix yn its M ethcc 



Reference 



rax i mum 

\~c Id i nc 1 1 me 



■.s M. eculis uc/g acid digestion E 3 A (1979) 6 months 
tissue AA/hot vacor 206.5/206.3 



<~H 



Cr 



_Cu 
Pb 
Mi 



ace digestion Stc. Met noes 

AA/coid vaocr I6tn ea. 2C2f 

acid diaestion E D A (1979) 

AA/flame 213.1 



218.1 
220.1 
239.1 



249.1 



289.1 



8. Project fiscal information 
Table 2. Est imated "Project Costs 



Total * samDies = 60 



Total cos.t for analysis 
@ Sl70.00/samole 



Total Project Cost = $10,200.00 



9. Schedule of Tasks and Projects 
(to be determined by DECE) 



94 



10. Project Organization and ResDonsibi I i ty 

Mr. W. Leign Bridges (Massachusetts Division of Marine Fisheries, 
Eoston, MA 02202, teleohone [6171 727-3194) will be the principal 
invesriaator for this project. He will be resDonsible to E?A for the 
t i me I y comD let ion of the project ana will have over a I I resDons ibi I i ty 
for data interprerat i on as well as preparation ana suomissicn or 
reoorts to E?A. 

Mr. Bridges will be assisted by Dr. Jack P. Schwartz (Division of 
Marine Fisheries. Cat Cove Marine Laooratory. Salem, MA 01S70, 
telecncne [617] 727-3958) as laboratory analysis leader. Dr. 
Scnwartz will be resocnsible for the orocessmc or all samoies 
rece:vec from DECE including duality control/cuai i tv assurance. 
anal vt i ca I- procedures, and ca:a storace and analysis. 



11. Data Quality Reduirements and Assessments 

Accuracy will be measured as oerdent recovery or an EPA stanaard 
reference material analyzed with each batch. Corrections will be made 
for badkgrouna levels. Average laooratorv recoveries will be 
maintained in the range of 80-120%. Unsptkea blanks will accompany 
every batch as a further measure of accuracy. 

Precision will be measured as the relative standard deviation of 
triplicate analyses perrormed uoon 10% of the samoLes in eacn batcn. 
Instrumental preci s ion wi I I be monitored through the use of triplicate 
readings on the transition metals oigestate or througn' tr ipl icate 
readings of oalibration standards 7 for arsenic and mercury. Should 
results vary by more than 10% readinas will be repeated. 

Completeness will be measured as the percentage of total samples 
received that were completely analyzed. We expect to achieve 100% 
completeness of a I -I analyses. 



12. Sampling and Analytical Procedures 

Elgi.d-Samp.Mng 

(to be determined by DEQE) 

Anaj_y_tj_caJ Pr.ocedu.res 

Arsenic analyses will be performed according to U.S. EPA method 
206.5/206.3. Mercury analyses will be performed according to Stanoarb 
Methods (16th ed.) 303F for the Examination of Water and Wastewater. 
Analysis for cadmium, dhromium, copper, lead, nickel, and zind will be 
performed using EPA methods 213.1, 218.1, 220.1, 239.1, 249.1, and 
289.1, respectively. All methods wi I I compliment EP A methods muse 
by DEQE. Concentrations of arsenic will be determined bv atomic 
aosorption hot vaoor technique. Mercury concentrations will be 



95 



determined by atomic adsorption cold vaoor techniaue. Transition 
metal concentrations will be determined by atomic aPsorption flame 
technigues. Analyses will be performed using a Perkin-Elmer Mocel 
3020E atomic absorDtion soect roDnotometer . Arsenic and mercury 
analyses will also use a Perkin-Elmer Mccel PHB 10 mercury hydride 
system. Samples will be ccmcared to external stanaarcs suitaoie for 
the metal being analyzed. 



12. Same I e Custodv Procedures. 

Hcmcqen i zee mussel samoles will be sniDDea frozen in polyethylene Pags 
dv DEGE cersonnei accompanied Pv an analysis recues: form (figure "). 
Laooratory cersonnei wi I l taxe custccv or ail sample materia: wmen 
wiil pe assigned laooratory tracking numoers (loggec-in) ana loekee in 
freezers. Due to a lack of space there are no plans to aremve 
samples. Any samo I e material remaining after the completion of all 
analyses will Pe made availaPle to DEQE. Mussel samples that are not 
frozen uDon oelivery will not Pe taken into custoay and returned to 
DEGE with the shipper. 



14. CaliPration Procedures and Preventive Maintenance 

The atomic aosprption soect roDnotometer will be calibrated through the 
use of external standards (certified atomic absorption grade standards 
ootained from Fisher Scientific Company). CaliPration of the 
instrument will occur at the Peginning of every sampling run and will 
be checked every ten samples and 'the end of every sampling run. 
Routine maintenance performed at the time of a run will be noted in 
the laboratory notebook. The instrument is covered by a maintenance 
contract with Perkin-Elmer. Any breakdowns will be promptly 
rerpai red. 



i«; 



Documentation, Data Reduction, and Reporting 



A. All raw data generated during laPoratory analysis will be kept in 
a permanently bound nctepook. A permanently Pound noteoook will be 
kept of all auality control tests conaucted at the laPoratory. Data 
printouts will be kept on file and avai ladle for inspection. 



16. Data Val idat ion 

All data produced Py the laboratory will pe suPiect to a 10055 check 
for errors in transcription and calculation Py the Senior Chemist. Dr 
Nina M. Duston, and the Laooratory Analysis Leader, Or. Jack P. 
Schwartz. The Principal Investigator, Mr. W. Leigh Bridges, will 
look at all logoooks and notepooks to ensure that reauirements are 
met. Data wnicn do not meet the SDecified auality reauirements will 
not Pe mcluaed in the report. Analytical reports will be signed by 
the Senior Chemist or Laooratory Analysis Leaaer Pefore being 
released. 



96 



17. Performance and System Audits 

Performance will be monitored through EPA water Pollution Laboratory 
Performance Evaluation Studies which provides for routine 
intercal ibrat ion with U.S. EPA every six months. 



18. Corrective Action 

Meetincs between all laooratory personnel and the Principal 
Investigator of the stuay will beheld at the comDietion of eacn 
samoie batch. Proolems will be identified as the stucv progresses. 
When corrective action is recuirea it will oe taken immec lately anc 
no zee m the appropriate laboratory ncteocoK. 



19. Reports 

The reports generated during this study are as follows: 

A. Quality assurance project plan, due May 29. 1987. This report 
will inciuae the objectives, scope, methods, and products associated 
wi th th i s study. 

8. At the completion of analyses of each samoie batch a report will 
be forwarded to the Principal Investigator for transmittal to 
appropriate U.S. EPA and DEQE personnel. This report will be 
completed before the next batch of samples is received. 



97 



DIVISION OF MARINE FISHERIES 

Laboratory Methodology 
Wet tissue Digestion Procedure for Trace Metals Analysis 

Chemicals 

1. HNO3 70.0 - 71.0% n Baker Instra-Analyzed Reagent for Trace Metal Analysis. 

2. H 2 2 , 30% Baker Analyzed Reagent. 

a. Weigh approximately 10 grams of blended tissue sample in a preweighed 
or tared tall form beaker (200 ml). Record sample wet weight to 
nearest 0.01 grams. 

b. Add 10 ml concentrated HN0 3 to sample in the tall form beaker. 
Cover with a watch glass and let sit overnight (15 to 16 hours) in 
ventilated fume hood to cold digest. 

c. Place covered samples on a steam bath until almost all tissue is 
digested. At this time spike the appropriate quality control samples 
with a standard spike solution containing concentrations as listed 
below for the particular species being digested. 



ialyte 


Finf ish 


Lobster 


Shellfish 




ppm 


ppm 


ppm 


Pb 


4.0 


4.0 


4.0 


Zn 


10.0 


50.0 


20.0 


Cu 


10.0 


50.0 


2.0 


Cr 


1.0 


1.0 


1.0 


Cd 


0.5 


0.5 


0.5 



(All standard solutions made in 2% V/V HN0 3 ) 

Use of these standard spike solutions will result in the enrichment 
values listed below for the final 50 mL volume of spiked digestate. 



alyte 


Finf ish 


Lobster 


Shellfish 




ppm 


ppm 


ppm 


Cd 


0.05 


0.05 


0.05 


Cr 


0.10 


0.10 


0.10 


Cu 


1.00 


5.00 


0.20 


Pb 


0.40 


0.40 


0.40 


Zn 


1.00 


5.00 


2.00 



4. Reflux the samples for 2 hours. 

5. Remove watch glass after 2 hours of refluxing and evaporate sample 
to near dryness. 



99 



6. Once all samples are evaporated to near dryness and are at room 
temperature, add 10 ml concentrated HN0 3 and 10 ml of 30% H 2 2 to each 
sample. Cover beaker with watch glass and let sit overnight (15 to 
16 hours) . 

7. Place covered samples on cold stream bath and slowly bring up to 
temperature. (Watch for violent reactions.) Reflux for 2 hours on 
steam bath. 

8. Remove watch glass and evaporate to near dryness. 

9. Add approximately 20 ml of a 2% v/v hot HNO3 solution to beaker and 
let heat for 5 minutes on steam bath. 

10. Remove beaker from steam bath, wipe off any moisture on the outside 
of beaker and filter the sample using a glass filter funnel with a 
reeve Angel 802 12.5 cm fluted filter paper or equivalent. Collect 
filtrate in 50 ml volumetric flask. Rinse beaker with two aliquots 
of 5-10 ml hot 2% v/v HN0 3 to remove as much yellow coloring as 
possible from the filter paper. Remove filter paper and rinse glass 
funnel with hot 2% HN0 3 taking care not to go over the 50 ml mark. 

11. Q.S. to 50 ml with 2% v/v HN0 3 and transfer to sample containers. 

12. Sample digestate is then analysed for metals on a Perkin Elmer AAS 
3030B according to the manufacturer's specifications. 



100 



Mercury Digestion Method 

Chemicals 

1. HN0 3 , 70.0-71.0%, "Baker Instra-Analyzed" Reagent for Trace Metal Analysis 

2. H 2 S0 4 , 95.0-98.0%, "Baker Instra-Analyzed" Reagent for Trace Metal Analysis 

3. KMn0 4 , "Baker Instra-Analyzed" Reagent for Hg Determination 

4. K2S 2 8 , "Baker Instra-Analyzed" Reagent for Hg Determination 
Solutions Needed 

1. 5% Potassium permanganate solution: Dissolve 25 g KMn0 4 in deionized 
distilled water and dilute to 500 ml. 

2. 5% Potassium persulfate solution: Dissolve 25 g K2S 2 8 in deionized 
distilled water and dilute to 500 ml 

Procedure for Shellfish Tissue Digestion 



1. Weigh approximately 2 grams of blended sample, to the nearest 0.1 mg, in 
a pre-weighed or tared 125 ml Erlenmeyer reaction flash. 

2. Add 7.0 ml cone. H 2 S0 4 and 3.0 ml HN0 3 to each flask and place in a 70 °C 
water bath. 

3. Remove samples to be spiked from water bath when a colored liquid with no 
visible tissue has formed. Spike appropriate Q.C. samples with 1.0 ml of 
100 ng/ml Hg. This will yield 50 ng Hg enrichment in final sample (refer 
to Step 8). Return samples to water bath. 

4. Samples should remain in the water bath for four (4) hours. 

5. Remove samples from water bath. Allow to cool to room temperature. Add 
5.0 mL deionized distilled water to the samples to cause precipitation of 
waxy digestion products and decrease the acidity of the sample solutions. 

6. Filter samples through VWR grade 615 9 cm or equivalent filter paper into 
a stoppered glass 25.0 mL graduated cylinder to remove the waxy 
precipitate. Rinse the sample flask twice with small amount of 20% v/ 
HNO3. Rinse filter paper with small amount of 20% HN0 3 taking care not to 
exceed 25.0 mL of liquid in cylinder. 

7. Q.S. to 25.0 mL with 20% HN0 3 . Stopper cylinder and shake well. 



101 



8. Using acid washed disposable 9 inch Pasteur pipets, divide sample into two 
equal portions and place in two clean 125 mL Erlenmeyer flasks. Rinse 
cylinder with two 2.5 mL portions of 20% HN0 3 solution, divide rinses 
equally between the two sample flasks. 

9. Ice samples. 

10. Add 10 mL KMn0 4 solution to each flask and let stand 15 minutes in ice 
bath. 

11. Add 8 mL K2S 2 O g solution to each flask while still in the ice bath. 

12. Add 0.5 - 1.5 g of KMn0 4 crystals as needed to keep the solutions purple. 
Remove from ice bath. Samples are left overnight to digest or are placed 
in a 70°C water bath for 2 to 4 hours. Please note, solutions must remain 
purple until analysis. Analysis must be with 24 hours. 



Washing Procedure for All Labware Used for Metals Analysis 



1. A 12 hour presoak is used if glassware has an organic/waxy film. The 
presoak solution is made from Terg-A-Zyme (as instructions indicate on the 
carton) . 

2. Wash with soap (Liquinox) and tap water, rinse well with tap water. 

3. Rinse thoroughly with 1:1 HN0 3 followed by 1:1 H 2 S0 4 (twice). A squeeze 
bottle is used to deliver the rinse. 

4. Rinse thoroughly with deionized distilled water at least three times. The 
deionized distilled water should have a resistance of 2Mohm or higher. 

5. Air dry or place in oven to dry. 

6. Store clean labware in assigned areas, covering with parafilm or glass 
stoppers, whichever is appropriate. 



102 







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